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Laboratory Techniques in
Genetic Engineering
Dr. H.K. Garg
Professor
Department of Biotechnology
Institute for Excellence in Higher Education, Bhopal
1
Contents
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
RNA Purification from Blood
Affinity Purification of Total RNA
Gene Isolation
Isolation of Cellular DNA
3.1
DNA Extraction from Strawberry Plant
3.2
Isolation of DNA from Banana
3.3
Isolation of Genomic DNA from Plant Source by
CTAB Method
3.4
Isolation of Chromosomal DNA by Lysozyme
Method
3.5
Isolation of Plasmid DNA from Bacteria
3.6
DNA Extraction: Organic Method
3.7
Silica Absorption Method
3.8
Inorganic Method
3.9
Chelex Method
3.10
Differential Method
Qualification of Nucleic Acids
4.1
Spectrophotometry
4.2
Characterization of DNA by Spectrophotometric
Assay and Melting Temperature (Tm)
4.3
Fast Technology Analysis
Recombinant DNA Technology
5.1
Designing DNA Probes
Bergs Terminal Transferase - Boyer Cohen Chang
Experiment
Preparation of Competent Cells for Efficient DNA Uptake
in E. coli
Bacterial Transformation in vitro Using Electroporation
8.1
CaCl2 mediated Transformation
Agarose Gel Electrophoresis
Protein Analysis by SDS-PAGE
Blue-White Colony Selection employing X-Gal / IPTG
Genomes and Interrelatedness
In situ Hybridization
Amplification of DNA using Polymerase Chain Reaction
05
09
13
22
24
26
28
31
34
36
39
41
44
46
48
50
54
60
64
66
69
72
77
80
83
88
92
96
101
110
2
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
35.
36.
14.1
Real-time PCR
14.1.1 Optimization of Annealing Temperature
14.1.2 Reverse Transcription PCR | RT-PCR
Digestion of DNA with RE
15.1
Digestion of DNA with RE in Bacteriophage
Southern Blotting of DNA
Northern Blotting of RNA
Western Blotting
Dot Blot
Immunoprecipitation
Sanger Sequencing
Maxam-Gilbert Sequencing
Pyrosequencing
Multiplex DNA Sequencing
Automated Sequencing
Construction of Molecular Maps
Restriction Mapping
Molecular Markers
28.1
Restriction Fragment Length Polymorphism
28.2
Random Amplified Polymorphic DNA
28.3
Amplified Fragment Length Polymorphism
28.4
DNA Chip and Microarrays
Genomic Library
cDNA Library
DNA Fingerprinting
Genetic Selection and Screening Method
32.1
Use of Chromogenic Substrates
32.2
Antibiotic Sensitivity Test for Detection of
Recombinants
32.3
Insertional Inactivation
32.4
Complementation of Defined Mutations
Protein Engineering
33.1
Rational Design
Stem Cell Therapy
Reverse Genetics
Transgenic Technology
36.1
Transgenic Plants
116
117
123
130
133
140
143
146
150
154
157
161
164
168
171
174
177
180
180
182
184
186
189
192
196
199
199
203
206
209
213
213
217
220
223
223
3
36.2
Transgenic Animals
Buffers and Reagents
Some Common Lexes
References
224
227
231
239
4
RNA Purification from Blood
RNA extraction is a technique used to isolate RNA from biological
samples. This is a challenging process due to the presence of
ribonuclease enzymes, which can quickly break down RNA. The most
common method used is TRIzol extraction, which starts with one full
Yellow-Top (type A) BD Vacutainer tube of human blood, roughly
equal to 8 milliliters, to yield around 30 micrograms of RNA. It's
essential to use RNAse-free tubes and solutions and maintain a clean
environment as RNA is highly sensitive to RNases, and autoclaving
does not inactivate them.
Principle
The TRIzol method
isothiocyanate,
a
is based on
powerful
protein
the use of
denaturant,
guanidinium
and
acidic
phenol/chloroform to separate RNA from the other components in
the sample. Keep the pH low so that only RNA will be separated in
the aqueous phase otherwise, at neutral pH, DNA will be separated.
Stock
1.
10x RBC Lysis Buffer
10.0 g KHCO3
2.0 ml 0.5 M EDTA
89.9 g NH4Cl
•
In a large container, dissolve 10.0 g of KHCO3, 2.0 ml of 0.5
M EDTA, and 89.9 g of NH4Cl in approximately 800 ml of
ddHO.
•
Adjust the pH of the solution to 7.3 by adding or removing
ddHO, as necessary.
5
•
Mix the solution thoroughly.
•
Transfer the solution to a tightly closed bottle and store at
2-8°C for up to 6 months.
2.
1x RBC Lysis Buffer
•
Take the 10x stock solution and dilute it 1:10 with ddHO.
•
Store the solution at room temperature up to 1 week.
3.
TRIzol Reagent or RNA STAT-60 Reagent:
Purchase either TRIzol Reagent (Invitrogen Life Technologies,
Cat No. 15596018) or RNA STAT-60 Reagent (Tel-Test, Cat
No. CS-111).
4.
Other Reagents
Phosphate Buffered Saline (PBS)
Isopropanol (2-propanol)
Ethanol
RNAse-free water
RNAse-Away (a cleaning solution to neutralize RNAses on
laboratory equipment).
Procedure
1.
Take a blood sample and transfer it to a 50 ml
polypropylene conical centrifuge tube. Adjust the volume of
the sample to 45 ml by adding RBC Lysis Buffer.
2.
Let the sample stand at room temperature for 10 minutes.
3.
Pellet the cells at 600 xg for 10 minutes using a centrifuge.
Remove the supernatant and gently resuspend the pellet in
1 ml of RBC Lysis Buffer.
4.
Transfer the resuspended pellet to a 1.5 ml microcentrifuge
tube.
6
5.
Pellet the cells for 2 minutes at room temperature using a
microcentrifuge. Remove the supernatant and resuspend the
pellet in 1 ml of sterile DPBS.
6.
Pellet the cells again and remove the supernatant.
7.
Add 1200 μl of TRIzol solution to each tube and resuspend
the cells. Add 0.2 ml of Chloroform (CHCl3) and vortex each
tube for 15 seconds.
8.
Centrifuge the samples at 13,000 rpm for 10 minutes at 4°C.
Transfer the upper phase to a clean microcentrifuge tube.
9.
Remove ~20% of the upper phase for future collection of
micro-RNA. Add an equal volume of cold isopropanol to the
remaining upper phase and mix by inverting.
10.
Store the samples in a -20°C freezer for precipitation.
11.
Centrifuge the samples again at 13,000 rpm for 10 minutes
at 4°C. Remove the supernatant and rinse the pellet with 0.5
ml of ice-cold 75% ethanol.
12.
Centrifuge the samples again. Remove the supernatant
along with any remaining liquid in the bottom of the tube
with a pipette.
13.
Allow the pellet to dry for 5 to 10 minutes. Dissolve the RNA
pellet by adding 20 μl of RNAse-free H2O to each sample.
14.
Measure the RNA concentration within 2 hours of elution
and store at -80°C.
Note: The stock solution of RBC Lysis Buffer is stable for 1 week at
room temperature and the RNA solution is stable for 6 months at 2–
8°C in a tightly closed bottle.
Advantages
TRIZol or Tri is a popular reagent used for RNA extraction. The
advantages of using this reagent include the combination of phenol
and guanidine isothiocyanate, which provides benefits from both
7
reagents. This combination effectively removes protein and DNA, but
the success of the extraction largely depends on the user's pipetting
skills. Also, the RNA is protected during the extraction process.
Disadvantages
However, there are also some disadvantages of using TRIzol or Tri.
The extraction process is heavily dependent on the user's pipetting
skills and any disturbance to the phases can result in contamination.
Furthermore, phenol and chloroform are harmful reagents that
should be handled with caution, such as under a hood. Finally, this
method of RNA isolation may not be suitable for some samples such
as low-input or degraded RNA.
8
Affinity Purification of Total RNA
The Affinity purification of total RNA involves isolating high-quality
mRNA for further use in molecular biology techniques. The principle
behind this technique is the affinity selection of polyadenylated
mRNA using oligodeoxythymidylate [Oligo (dT)]. In simple terms, this
means that the process uses a specific type of molecule,
oligodeoxythymidylate, to selectively target and purify the mRNA
that has a polyadenylated tail. This allows for the isolation of highquality mRNA for further use in molecular biology techniques.
The first step in this process is to extract the total RNA from the
sample of interest. This RNA contains both mRNA and non-coding
RNA. Thereafter, the extracted RNA is mixed with oligodeoxythymidylate, which will specifically bind to the polyadenylated
mRNA. The mixture is then passed through a column that separates
the mRNA bound to the oligodeoxythymidylate from the rest of the
RNA. Finally, the column is washed to remove any contaminants and
the purified mRNA is eluted, or released, from the column. This highquality mRNA is then ready for use in molecular biology techniques,
such as cDNA library construction.
Stock
To perform this procedure, several materials are required. It's crucial
that all materials used in this process are sterile and of molecular
biology grade, including:
RNase-free water
SDS
Oligodeoxythymidylate-cellulose [oligo (dT)]
RNase-free glass wool
9
Pasteur pipettes
5 M NaCl
3 M Sodium acetate pH-6.0
Absolute alcohol, 70% ethanol
Loading buffer
Elution buffer
Recycling buffer.
All Tris-containing solutions should be prepared using RNase-free
water and autoclaved, while all other solutions should be treated
with diethyl pyrocarbonate (DEPC) and autoclaved, but for Tris,
which inactivates DEPC.
To prepare RNase-free water, add 0.1% DEPC to water, let it stand
overnight at 37°C, and then autoclave it to destroy residual DEPC
activity. SDS should be weighed in a fume hood, as it is dangerous, if
inhaled. A 10% stock solution is usually prepared, but it is unstable if
autoclaved, so it should be heated at 65°C for 2 hours to destroy any
residual RNase activity. Oligo (dT) cellulose should be purchased
commercially and suspended in loading buffer at a concentration of
5 mg/1 ml. It should be stored either dry at 4°C or in suspended
condition in loading buffer at -20°C. The glass wool and Pasteur
pipettes should be wrapped in aluminum foil and baked at 200°C for
2 to 4 hours to remove any RNase activity. The rest of the materials
should be stored as follows:
5 M NaCl (Store at room temperature).
3 M Sodium acetate pH-6 (Store at room temperature).
Absolute alcohol (Store at -20°C).
70% ethanol (Prepare this solution using DEPC-treated
water. Store at 4°C).
Loading buffer (0.5 M NaCl in 0.5% SDS, 1 mM EDTA, 10 mM
Tris-HCl - pH 7.5 Store at room temperature).
10
Elution buffer (1 mM EDTA, 10 mM Tris-HCl - pH 7.5. The
buffer can be stored at room temperature but should be
preheated to 65°C prior to use).
Recycling buffer (0.1 M NaOH, which should be prepared
immediately before use and used fresh).
Note that this method is dependent on the purity of the samples,
and it may not work on low-input or degraded RNA. Also, when
handling DEPC, gloves should be worn, as hands are a major source
of RNase activity, and DEPC is a carcinogen that should be handled
in a fume hood with extreme care.
Procedure
Step 1: Preparing an Oligo (dT) Column:
1.
Take a syringe and remove the plunger.
2.
Fill the base of the syringe with glass wool.
3.
Take a sterile, RNase-free Pasteur pipette and add the
oligo(dT) cellulose on top of the glass wool.
4.
The oligo(dT) cellulose will form a column above the glass
wool.
5.
Oligo (dT) column is now ready for use.
Step 2: Isolating Poly (A+) RNA
1.
Resuspend the RNA pellet in loading buffer.
2.
Heat the RNA pellet to denature it.
3.
Load the denatured RNA onto the oligo (dT) column.
4.
Wash the column to remove any unbound RNA.
5.
Add elution buffer to the column to recover the bound poly
(A+) mRNA.
6.
Precipitate and wash the mRNA to obtain pure and intact
mRNA.
11
Advantages
1.
Elimination of organic solvents.
2.
Compatible with a variety of sample types.
3.
DNase treatment to eliminate contaminating genomic DNA.
Disadvantages
1.
May be less efficient in obtaining RNA.
2.
More expensive compared to other methods.
12
Gene Isolation
Gene isolation is a process in molecular biology and genetics that
involves separating a single gene from its surrounding DNA
sequence in the genome. The goal of gene isolation is to obtain a
pure, functional copy of a gene, which can then be studied,
manipulated, or expressed in vitro. The process of gene isolation can
involve a variety of techniques, including restriction enzyme analysis,
PCR, and hybridization, and can be used to study the function of
specific genes or to develop new therapies for genetic diseases.
DNA Extraction and Purification
DNA is a crucial element in molecular biology and can be found in
various sources including human tissues, blood, hair, leaves, bacteria,
insects, and more. DNA extraction is a process of taking DNA from
cells or tissues of interest. This technique is the first step in the study
of specific DNA sequences, genomic structure, DNA fingerprinting,
restriction fragment length polymorphism (RFLP), and PCR analysis.
DNA extraction is done through different methods such as
mechanical methods where the cells are blended to release the DNA,
chemical methods where detergents or enzymes are used to break
down the cell membrane or physical methods where centrifugation
is used to separate the DNA. Apart from these, there are various
methods that can be used to perform DNA extraction, such as
organic extraction, salting out, magnetic separation and silica-based
technology. The choice of method depends on factors such as tissue
type, DNA concentration, sample number, safety, and cost.
13
After extraction, the DNA is purified to get rid of any contaminants
that may still be present. This is done through methods such as
ethanol precipitation, column-based purification, or size-exclusion
chromatography. The final purified DNA is then stored in a buffer
solution at low temperature for future use.
There are various techniques used for DNA extraction, but all of
them include the following fundamental processes:
1.
Cell lysis: This is the breakdown of the cellular structure,
which releases long strands of DNA. Depending on the
source, cell lysis can be done chemically, physically, or a
combination of both. For example, plant and bacterial
samples' cell walls are broken by applying physical force,
while chemical agents such as lysozyme, EDTA, and
detergents are used for lysis in other sources.
2.
Elimination of membrane lipids: After lysis, DNA is processed
to remove further contaminants before the membrane lipids
are extracted. This is done by washing the DNA.
3.
Protein denaturation and removal: Proteins can interfere
with molecular biology experiments, so they are denatured
and removed using an enzyme called protease.
4.
Removal of additional cellular elements: Through repeated
washing
processes,
other
biological
components
are
separated from the DNA.
5.
Denaturation and removal of RNA: RNA is a significant DNA
contaminant, so it is denatured and removed using an
enzyme called RNase.
14
6.
DNA elution and storage: The purified DNA is eluted in an
alkaline buffer solution or in double distilled water and
stored at -20°C for future use.
Stock
Chemical
Ethylene diamine tetra acetate (EDTA), NaOH, Tris-HCl, sucrose,
MgCl2, Triton X100, Sodium dodecyl sulphate (SDS), NaCl, Sodium
perchlorate, TE buffer or double distilled water, cold chloroform, cold
ethanol.
Preparation of solutions
1.
0.5 M EDTA, pH 8.0
Add 146.1 g of anhydrous EDTA to 800 ml of distilled water.
Adjust pH to 8.0 with NaOH (about 20 g). Make up the
volume to 1 L with distilled water.
2.
1 M Tris-HCl, pH 7.6
Dissolve 121.1 g of Tris base in 800 ml of distilled water.
Adjust pH with concentrated HCl (about 60 ml). Make up the
volume to 1 L with distilled water.
3.
Reagent A (Red Blood Cell Lysis Solution)
0.01M Tris-HCl (pH 7.4), 320 mM Sucrose, 5 mM MgCl2, and
1% Triton X100.
Add 10 ml of 1 M Tris to 109.54 g of sucrose, 0.47 g MgCl2
and 10 ml Triton X100 to 800 of distilled water. Adjust pH to
8.0; make up the volume to 1 L with distilled water.
4.
Reagent B (White blood Cell Lysis Solution)
15
0.4 M Tris-HCl, 150 mM NaCl, 0.06 M EDTA, 1% SDS, pH 8.0.
Take 400 ml of 1 M Tris (pH 7.6), 120 ml of 0.5 M EDTA (pH
8.0), 8.75 g of NaCl, adjust pH to 8.0 with NaOH. Make up
the volume to 1 L with distilled water. Autoclave at 15 psi for
15 min. After autoclaving the mixture, add 10 g of SDS.
Procedure
1.
Obtain 3 ml of whole blood and place it in a 15-ml falcon
tube.
2.
Add 12 ml of reagent A to the tube and mix for 4 minutes
using a rolling or rotating blood mixer at room temperature.
3.
Centrifuge the mixture at 3000g for 5 minutes at room
temperature.
4.
Discard the supernatant without disturbing the cell pellet
and remove any remaining moisture by blotting onto tissue
paper.
5.
Add reagent B and 250 μL of 5 M NaCl to the tube and mix
by inverting several times.
6.
Place the mixture in a water bath at 65°C for 15 to 20
minutes.
7.
Add 2 ml of ice-cold chloroform and mix on a shaker for 20
minutes.
8.
Centrifuge the mixture at 2400g for 2 minutes.
9.
Transfer the upper phase to a clean falcon tube using a
sterile pipette.
10.
Add 2 to 3 ml of ice-cold ethanol to the tube and gently
invert to allow DNA to precipitate. If necessary, add more
ethanol.
11.
Using a clean Pasteur pipette, spool the DNA onto the
hooked end and transfer to a 1.5-ml microcentrifuge tube.
16
12.
Centrifuge the microcentrifuge tube at 6000 rpm for 5
minutes.
13.
Gently remove the supernatant (ethanol layer) without
disrupting the DNA pellet and leave to dry.
14.
Resuspend the DNA in 200 μL of TE buffer or doubled distal
water and label.
15.
Determine the yield and purity of the extracted nucleic acid
if necessary. The results should show a cloudy precipitation,
representing the isolated genomic DNA, visible to the naked
eye.
Alternative Method
Step 1: Lysis
Add lysis buffer to the blood sample
The lysis buffer will break open the red blood cells and
release the DNA into solution
Step 2: Precipitation
Add alcohol (such as ethanol or isopropanol) to the lysed
blood sample
The DNA will precipitate out of solution and can be collected
by centrifugation
Step 3: Wash
Wash the DNA pellet with a buffer to remove remaining
contaminants
Step 4: Resuspension
Resuspend the purified DNA in a buffer of your choice
17
Step 5: Quantification
Quantify the purified DNA using spectrophotometry or
fluorometry.
Alternate method to extract genomic DNA from lizard
Stock
1.
SE
Sodium chloride (75 mM)1M
37.5 ml
EDTA (25 mM) 500 Mm 25.0 ml
Distilled water
2.
437.5 ml
Sodium acetate 3M
Sodium acetate 24.6 gm
Distilled water
100.0 ml
(adjust pH to 5.2 with glacial acetic acid)
3.
TEN
Tris (pH 8.0) (10 mM)
1M 5.0 ml
Sodium chloride (100 mM) 1 M 50 ml
EDTA (pH 8.0) (25 mM) 500 mM 25 ml
Distilled water
4.
420 ml
TE
Tris (pH 8.0) (10 mM)
1 M 1ml
EDTA (1 mM) 500 mM
0.2 ml
Distilled water
98.8 ml
5.
SDS 10% and Proteinase K (10 mg/ml)
6.
Distilled and buffered saturated phenol
7.
Phenol–Chloroform-Isoamyl alcohol (25:24:1)
8.
Absolute alcohol
9.
Ethanol 70%, prechilled at - 20oC
18
Laboratory ware
1.
Homogenizer
2.
Micropipettes
3.
Pipette tips
4.
Centrifuge tubes
5.
Eppendorf tubes
6.
Cheese clothe
Procedure
1.
Homogenize a Calotes embryo at 4°C in SE (salt-EDTA
buffer).
2.
Centrifuge the homogenized sample at 4000 rpm for 10
minutes at 4°C, then discard the excess liquid.
3.
Redissolve the remaining particle in 20 ml of TEN (TrisEDTA-NaCl) buffer at room temperature.
4.
Add SDS (sodium dodecyl sulfate) to the mixture to a final
concentration
of
1%
and
proteinase
K
to
a
final
concentration of 100 ug/ml.
5.
Incubate the mixture at 37°C for an overnight period.
6.
Add an equal amount of phenol to the mixture, stir in a
vortex mixer for 15 minutes, then centrifuge for 10 minutes
at 12000 rpm.
7.
Transfer the aqueous phase from the previous step to a new
tube, and add an equal volume of phenol, chloroform, and
isoamyl alcohol. Centrifuge and discard the organic phase.
8.
9.
Repeat the above step to further purify the DNA.
Add an equal volume of chloroform-isoamyl alcohol to the
aqueous phase, transfer the aqueous phase to a new tube
after centrifugation.
10.
Combine 1/30 volume of sodium acetate (3M) with 2
volumes of 100% ethanol and maintain at -20°C for 1 hour.
19
11.
Transfer the precipitated DNA to a new Eppendorf tube
containing 70% ethanol (chilled at -20°C).
12.
Centrifuge the tube briefly, extract as much of the remaining
alcohol as possible, lyophilize the DNA, and dissolve in TE
(Tris-EDTA) buffer.
Alternative Method
Step 1: Tissue Collection
Obtain fresh or frozen tissue samples from the lizard, such as
blood, muscle, or liver.
Store the tissue samples in a clean and sterile container.
Step 2: Lysis
Homogenize the tissue samples using a mechanical
Ensure that the tissue is fully lysed to release the cellular
homogenizer or a chemical lysis buffer.
contents, including the DNA.
Step 3: Cell Debris Removal
Centrifuge the lysate to separate the cell debris from the
DNA-containing supernatant.
Carefully pour off the supernatant, leaving behind the cell
debris.
Step 4: DNA Purification
Use a DNA purification kit or ethanol precipitation to purify
the DNA from the supernatant.
Follow the instructions provided with the DNA purification
kit or refer to the appropriate protocols for ethanol
precipitation.
20
Step 5: DNA Quality and Quantity
Check the quality and quantity of the purified DNA using
methods such as spectrophotometry, gel electrophoresis, or
PCR.
Ensure that the purified DNA is of sufficient quality and
quantity for your intended use.
Notes
The specific protocol may vary based on the source of the tissue and
the DNA purification kit used. To avoid cross-contamination, take
proper precautions and maintain sterile conditions throughout the
extraction process.
21
Isolation of Cellular DNA
Stock
Coconut endosperm
Sodium chloride
Sodium citrate
Mortar and pestle
Centrifuge tubes
Absolute alcohol
Procedure
1.
Grind 200mg of tissue (coconut endosperm, spleen, heart,
testis, or kidney) in a saline citrate solution (85ml 0.9%
sodium chloride solution and 15ml 0.5% sodium citrate
solution with pH 7.4).
2.
Transfer the homogenate to a centrifuge tube, bring the
volume to 10ml with saline citrate solution.
3.
Centrifuge at 3000 rpm for 8 minutes, discard the
supernatant.
4.
Re-homogenize the pellet with 5 ml saline citrate solution,
adjust the volume to 10 ml, repeat the centrifugation
process for 8 minutes and discard the supernatant.
5.
Suspend the pellet in 12% sodium chloride and centrifuge at
10,000 rpm for 15 minutes in a refrigerated centrifuge.
6.
Transfer the supernatant to a 30 ml test tube, add 2-3
volumes of absolute alcohol, mix gently.
7.
Collect the fibrous white DNA by winding it around a clean
glass rod.
22
8.
Transfer the DNA into a 1.5 ml Eppendorf tube, add 1 ml
70% alcohol, centrifuge at 1000 rpm for 5 minutes, discard
the supernatant.
9.
Dry the DNA pellet; dissolve in 2 ml distilled water, measure
optical
density
at
260
nm
wavelength
in
a
spectrophotometer.
23
DNA Extraction from Strawberry
Plant
The process of isolating pure genomic DNA from plant tissue is
crucial for studying plant genetics and making changes to plant
genes and metabolic pathways. This process is different from
extracting DNA from animal sources due to the presence of a tough
cellulose cell wall and a larger DNA molecule.
The goal of the isolation process is to obtain pure and high-quality
DNA without any cellular material or degradation. To achieve this,
the cell wall and cellular membranes need to be broken down
through either mechanical or non-mechanical methods. Mechanical
methods use physical force to open the cell wall, while nonmechanical methods use enzymes or chemicals along with physical
force to break down the cell wall components.
Once the cell wall is broken, the cell membrane forms small cracks,
which allows for the use of detergents to break down the cell
membrane. The DNA is then separated from the protein by using
isopropanol or ethanol. The final product is a clean DNA suspended
in a buffer or distilled water.
Stock
Chemical
1.
Strawberry
2.
Extraction solution
3.
96% cold ethanol or isopropanol
24
4.
TE buffer or double distilled water.
5.
Preparation of extraction solution
Add 100 ml detergent to 750 ml of distilled water and then add 11 g
NaCl. Make up the volume to 1 L with distilled water.
Equipment and Glassware
Microfuge centrifuge, electronic balance, razor blade, mortar and
pestle, cheese cloth, funnel, graduated cylinder 25 ml, beaker 50 ml,
test tube, centrifuge tube, Pasteur pipette, micropipette, and tips.
Procedure
1.
Remove the green leaves from the strawberry and weigh the
plant using a sensitive balance.
2.
Chop the plant into small pieces using a clean razor blade.
3.
Mix the chopped plant with the extraction solution for 5
minutes using a pestle.
4.
Pour the mixture through cheese cloth into a clean beaker.
5.
Pipette a small amount of the mixture into a test tube and
add cold ethanol or isopropanol.
6.
The DNA will appear as a clear white thread, spool it onto
the hooked end of a Pasteur pipette.
7.
Transfer the DNA to a centrifuge tube and spin it at 6000
rpm for 5 minutes.
8.
Remove the supernatant (ethanol layer) gently, leaving the
DNA pellets to dry.
9.
Suspend the DNA pellets in TE buffer or double distilled
water.
Result
Cloudy precipitation will be obtained, and it represents the isolated
DNA.
25
Isolation of DNA from Banana
Stock
Fleshy berry fruits like banana, grapes, etc.
Liquid soap
Distilled water
Salt (NaCl)
Ice-cold isopropyl alcohol (IPA)
Measuring spoons
Glass stirring rod
Test tubes
Glass beakers
Plastic cups
Strainer or coffee filter
Funnel
Principle
The DNA extraction protocol involves disrupting the cell wall, cell
membrane, and nuclear membrane of plant cells to release the DNA
into
solution,
followed
by
precipitation
and
removal
of
contaminating biomolecules.
Procedure
1.
Grind a piece of banana in a mortar and pestle to form a
mash or blend.
2.
Add 1/2 cup of distilled water to the banana mash and
transfer the mixture into a glass beaker.
26
3.
In a plastic cup or zip lock bag, mix 1 teaspoon of liquid
soap and 1 teaspoon of salt with 2 tablespoons of distilled
water. Stir gently until the salt and soap are dissolved.
4.
Add 2 tablespoons of the banana mash mixture to the cup
containing the salt and soap solution. Stir the mixture for 1015 minutes using a glass stirring rod.
5.
Filter the fruit mixture through a fine sieve or coffee filter.
6.
Chill a test tube of IPA by placing it in a beaker containing
ice cubes and water.
7.
Using a dropper, slowly add the filtrate to the chilled IPA in
the test tube.
8.
9.
Place the test tube undisturbed for 5-6 minutes.
The isolated DNA will appear as a white precipitate in the
alcohol layer.
10.
Gently spool out the DNA using a hook, bent paperclip, or
glass stirring rod.
Observation
The DNA will precipitate out into the alcohol layer, appearing as
white stringy mucus.
Result
The experiment yielded a good quantity of DNA, making it suitable
for use in biological experiments and biotechnology applications.
27
Isolation of Genomic DNA from
Plant Source by CTAB method
Stock
Young and tender leaves (Tulsi and Bryophyllum)
Mortar and pestle
Conical flask
Measuring cylinder
Distilled water
Ethidium Bromide
Ethanol
Micropipette
Centrifuge
Electrophoresis unit
CTAB buffer
Water bath
Eppendorf tube
CTAB buffer:
CTAB - 2 g
Tris HCl (1 M)10 ml
EDTA (0.5 M) 4 ml
NaCl (5 M) 28 ml
H2O 40 ml
PVP 40 1 g
28
Principle
The CTAB method uses a detergent to break open plant cells and
solubilize their contents. The DNA is then extracted from the cell
homogenate, with RNA removed by RNAase, and DNA precipitated
and washed in organic solvents before being redissolved in aqueous
solutions.
Procedure
1.
Take young and tender leaves, wash them with distilled
water, and grind them using a mortar and pestle.
2.
Transfer the powdered leaves into an Eppendorf tube and
add 50 ml of CTAB buffer.
3.
Incubate the plant homogenate for 15 minutes at 55°C in a
4.
After incubation, spin the tube at 12,000 rpm for 5 minutes
water bath.
at 4°C. Transfer supernatant to another Eppendorf tube and
discard the pellet.
5.
Add 250 ml of phenol: chloroform: isomyl alcohol (25: 24: 1)
and mix the solution.
6.
Spin the tube at 12,000 rpm for 1 minute at room
temperature. Transfer the aqueous phase to another tube.
7.
Add 500 ml of isopropanol and mix by inverting the
centrifuged tube at 10,000 rpm for 5 minutes at room
temperature. Discard the supernatant.
8.
Add 1 ml of 70% ethanol and mix with the pellet by inverting
it.
9.
Centrifuge the tube at 12,000 rpm for 1 minute at room
temperature.
10.
Discard the supernatant and let the pellet dry on ice for 15 30 minutes.
11.
Add distilled water to dissolve the DNA.
29
12.
Warm the DNA solution at 65°C for 20 minutes.
13.
Quantitate the DNA and check its purity.
14.
Run the DNA on a gel.
Results
DNA was successfully isolated from the leaves using the CTAB
method.
Precautions
Use soft leaves.
Avoid over-drying the DNA as it may become difficult to
suspend in TE.
Handle the water bath carefully.
Pre-chill the CTAB buffer and wash it in 70% ethanol before
use.
30
Isolation of Chromosomal DNA
from E. coli using the Lysozyme
Method
To isolate chromosomal DNA, mechanical barriers in bacterial cells
need to be disrupted, including the plasma membrane, cell wall, and
outer membrane in gram-negative bacteria. In this experiment, Trisbuffer containing EDTA is used to make the cells isotonic and
prevent them from bursting. Lysozyme treatment is used to attack
N-Acetalglucosamine residues of bacterial cell walls, making the
weakened cell wall porous and exposing periplasmic spaces. SDS
treatment is used to dissociate the cell membrane, followed by
treatment with phenol-chloroform to denature proteins and separate
aqueous organic phases. Chilled isopropanol is used to precipitate
DNA from the aqueous phase, and reprecipitation with 70% ethanol
is performed to eliminate divalent cations. The resulting DNA palate
is relatively pure and is suspended in Tris-EDTA buffer.
Stock
E. coli cells
Saline-EDTA
SDS (sodium dodecyl sulphate)
Chloroform: isoamyl alcohol
Ethanol
Phenol
Tris-HCl
NaCl
31
RNase
Chemical preparation
TE buffer: Dissolve Tris-HCl and EDTA together at their
respective molarities to create a single solution.
TGE buffer: Mix 25 mM Tris-HCl pH 8.0, 50 mM glucose, and
10 mM EDTA.
10% SDS of pH 6.8-7.2.
10 mg/ml RNase: Weigh 10 mg pancreatic ribonuclease and
dissolve in 1 ml of 100 mM Tris-HCl of pH 7.5 containing 150
mM NaCl. Boil in a water bath for 10 minutes.
Prepare 70% (v/v) ethanol in autoclaved double-distilled
water.
Chill isopropanol by refrigeration.
Lysozyme: Dispense 20 mg lysozyme in 5 ml TGE buffer and
keep on ice until use.
Solutions preparation:
Solution
A:
Saturated
phenol
with
0.1
g
of
α-
hydroxyquinoline and 100 ml melted phenol, followed by
transfer into a separating funnel to allow layers to form.
Remove
the
aqueous
layer
containing
Tris-HCl
and
impurities. Collect the phenol layer in an amber-colored
bottle.
Solution B: Mix chloroform and isoamyl alcohol in the ratio
24:1.
Solution C: Mix solution A and B in the ratio 1:1.
Procedure
1.
Grow E. coli cells in LB broth until they reach the log phase
(indicated by an absorbance of 0.4). Harvest 50 ml of
bacterial suspension by centrifuging at 10,000 rpm for 10
minutes. Gently discard the supernatant.
32
2.
Resuspend the pellet in 1 ml of TGE buffer. Spin the mixture
at 5,000 rpm for 5 minutes and carefully remove the
supernatant.
3.
Add 150 μL of lysozyme solution, vortex, and let it sit at
room temperature for 30 minutes.
4.
Slowly add 30 μL of RNase solution to the walls of the
Eppendorf tubes, and incubate them undisturbed at 37°C for
30 minutes.
5.
Slowly add 50 μL of 10% SDS to the walls of the tubes, mix
gently to prevent the formation of froth, and incubate at
37°C for 2 hours.
6.
Add 230 μL of solution C and vortex for 2 minutes to mix
well. Centrifuge the mixture at 10,000 rpm for 15 minutes at
4°C.
7.
Using a micropipette, carefully transfer the supernatant
aqueous phase to another tube. Add an equal volume of icecold isopropanol and mix well.
8.
Spin the mixture at 10,000 rpm at 4°C for 10 minutes and
carefully discard the supernatant.
9.
Add 0.5 ml of chilled 70% ethanol to the pellet to precipitate
the DNA. Recover the DNA by centrifuging at 10,000 rpm for
10 minutes and discard the supernatant. Dry the pellet in air.
10.
Dissolve the pellet in 50-100 μL of 10 mM TE buffer and
keep at 50°C for 5 minutes for better dissolution.
Electrophorese the isolated DNA on an agarose gel.
Result
The lysozyme method successfully isolated DNA from bacteria.
Precautions
Wear gloves while performing the experiment.
Handle phenol with care.
33
Isolation of Plasmid DNA from
Bacteria
Plasmids are circular DNA molecules found in bacteria that carry
genetic information separate from the chromosomal DNA. Their
isolation is significant for genetic engineering as plasmids can confer
resistance to various substances, act as vectors in recombinant DNA
technology, and play a role in bacteria's survival and adaptation.
In order to isolate plasmid DNA, bacteria are grown in a suitable
medium overnight or until reaching an optical density of 0.6-1.0 at
600 nm, and stored, if necessary, at 4°C. The bacteria are then lysed
with a lysis buffer, lysozyme, Tris-HCl, EDTA, and NaCl, and SDS. The
lysozyme degrades the bacterial cell wall, while the Tris-HCl, EDTA,
and NaCl neutralize the charge and protect the DNA from
degradation by nucleases. The SDS denatures the chromosomal DNA
and makes it more accessible to degradation by proteases.
The plasmid DNA can be extracted through phenol-chloroform
extraction and ethanol precipitation, followed by washing with 70%
ethanol and resuspending in TE buffer. The purified plasmid DNA
can be used for further experiments like restriction digestion.
Stock
1.
Bacterial culture
2.
Lysis buffer (such as lysozyme, Tris-HCl, EDTA, NaCl)
3.
Sodium dodecyl sulfate (SDS)
4.
Phenol-chloroform
34
5.
Isopropanol
Equipment
1.
Centrifuge
2.
Microcentrifuge tubes
Procedure
1.
Grow the bacterial culture in a suitable medium, such as
Luria-Bertani (LB) broth, overnight to reach an optimal
density of bacteria.
2.
Lyse the cells by adding a lysis buffer such as a solution of
NaOH, EDTA, and SDS and incubating the mixture at 65°C
for 10 minutes. This will break open the bacterial cell walls
and release the plasmid DNA into the solution.
3.
Neutralize the lysis solution by adding a neutralization
buffer, such as Tris-HCl, to restore the pH to 7.0.
4.
Centrifuge the lysis solution at high speed to separate the
bacterial debris (protein and other contaminants) from the
supernatant containing the plasmid DNA.
5.
Precipitate the plasmid DNA from the supernatant by adding
an equal volume of isopropanol. The isopropanol will cause
the plasmid DNA to form a dense band, which can be
collected and purified.
6.
Dialyze the plasmid DNA to remove the excess salts and
impurities.
7.
Precipitate the plasmid DNA from the dialysis solution by
adding ethanol and sodium acetate. The plasmid DNA will
form a pellet, which can be washed with 70% ethanol to
remove any remaining impurities.
8.
Dry the plasmid DNA pellet, and dissolve it in a suitable
buffer, such as TE buffer, for downstream applications.
35
DNA Extraction
Organic Method
The process of extracting DNA from a material involves several
important factors, including the efficiency of the extraction, the
amount of DNA obtained, the removal of impurities, and the quality
and purity of the DNA. One common technique for DNA extraction is
the organic method, also known as the phenol-chloroform method.
This method is particularly effective for extracting large amounts of
high molecular weight DNA, which was necessary for early DNA
fingerprinting techniques.
Principle
The principle behind this method is to mix a watery sample with a
solution of phenol and chloroform in equal parts. Upon mixing and
centrifugation, the mixture separates into two distinct layers - an
aqueous phase on top and an organic phase at the bottom. The
nucleic acids and other impurities stay in the aqueous phase, while
proteins move into the organic phase.
Stock (Reagents)
1.
Chloroform
2.
Ethanol
Ether (optional)
Nucleic acid solution to be purified
Phenol:Chloroform (1:1)
Tris EDTA(pH7.8) (optional)
3 M sodium acetate pH 5.2 or5 M ammonium acetate
36
100% ethanol
Equipment
1.
Automatic pipette fitted with a disposable tip
2.
Pipettes, large-bore (optional)
3.
Polypropylene tube
4.
Rotating wheel(optional)
Procedure
The step-wise method for the organic method is as follows:
1.
Add an equal volume of phenol: chloroform to the nucleic
acid sample. Mix the tube's contents to create an emulsion.
2.
Centrifuge the mixture for one minute at room temperature
at 80% of the tubes' maximum speed. Transfer the aqueous
phase to a new tube, discarding the organic phase and
interface.
3.
Repeat steps 1-4 until there are no longer any proteins
visible at the organic / aqueous phase interface.
4.
Repeat steps 2-4 with an equal volume of chloroform.
Measure the aqueous phase's volume in order to retrieve
DNA.
5.
Add 0.1 volume of pH 5.2 sodium acetate with a 0.3 M final
concentration.
6.
Add two to three litres of cold 100% ethanol. Keep at -20
degrees Celsius for longer than 20 minutes.
7.
Spin a microfuge for 10-15 minutes at its fastest speed.
Decant the supernatant carefully.
8.
Add 1 cc of 70% ethanol and mix. Spin quickly and decant
the supernatant with care.
37
9.
Dry the pellet with air or briefly vacuum. Resuspend the
pellet in the required volume of double-distilled water or
Tris EDTA buffer.
10.
Store and carry out quantification and intended usage
The approach discussed has various advantages, one being its
versatility in being used on different samples. However, it also has
some limitations, such as being a time-consuming process, being
susceptible to contamination and posing a risk to the scientist as it
involves handling hazardous substances.
38
Silica Absorption Method
The silica absorption method is a widely used process for purifying
DNA from different sources like blood, saliva, or plant tissue. This
process involves several steps to isolate the DNA from impurities like
proteins, lipids, and polysaccharides. Firstly, the sample is lysed using
a buffer containing detergents to release the DNA. Then, the lysate is
mixed with a solution containing silica particles and high salt
concentrations, (which cause the DNA to bind to the silica under
specific conditions, such as the use of specific salts and pH levels)
and the impurities remain in the solution. The silica-DNA complex is
then washed with a buffer containing high salt to remove impurities.
The purified DNA is finally eluted from the silica particles by reducing
the salt concentration. The purified DNA can then be checked for
quality and quantity.
There are variations of this method, such as using chaotropic agents
to enhance the efficiency of purification, and the choice of buffer,
salt concentration, and elution method can impact the purity and
yield of the purified DNA. The protocol may also need to be adjusted
for different types of samples, such as FFPE samples.
It's essential to be careful of cross-contamination and to take
appropriate measures to reduce the risk of contamination. Although
the exact mechanism of the silica DNA extraction method is not
entirely understood, it remains a widely used technique in many
laboratories.
39
Stock
To perform this technique, kits are available that include all
necessary materials.
Procedure
1.
Prepare the DNA sample for the extraction process.
2.
Run the sample through a microchannel.
3.
Allow the DNA to bind to the silica in the channel.
4.
Wash away any other molecules in the buffer solution.
5.
Purify the channel to remove any contaminants.
6.
Dry the silica.
7.
Extract the DNA by using either water or a buffer with low
salt concentration.
8.
Collect the DNA at the end of the channel.
Advantages
Quick
Dependable
High-quality DNA yield
Disadvantages
Expensive
Interference from certain sample sources like chewing gum.
40
Inorganic method
The non-organic method is a way to clean up DNA or RNA without
using any chemicals derived from organic materials. The key to this
method is adding a specific enzyme, Proteinase K, to the mix. This
enzyme helps to protect the DNA or RNA from other enzymes,
known as nucleases that can break down the nucleic acids during the
purification process.
Stock
Reagents
Digestion Buffer (10 mM NaCl, 10 mM TRIS (pH 8.0), 10 mM
EDTA (pH 8.0),
0.5%SDS
Proteinase K (20 mg/ml)
Sodium Acetate pH 5.2 (3M)
Ice-cold 98% ethanol
Ice-cold 70% ethanol
1X TE
Water
Tissue
Equipment
1.
Incubator
2.
Centrifuge
3.
Sterile 1.5-ml micro-centrifuge tubes
41
Procedure
The non-organic method includes:
Step 1: Tissue Digestion
1.
Obtain a clean micro-centrifuge tube and add 1.5 ml of the
digestion buffer.
2.
Calculate the appropriate amount of proteinase K to add to
the buffer based on the formula: 5 μl of proteinase K for
every ml of digestion buffer.
3.
Homogenize the tissue in the solution by mixing.
4.
Incubate the mixture for 1-12 hours at 55°C. This can be
done overnight.
5.
Vortex the mixture for a short period of time.
6.
Centrifuge the mixture at high speed for two minutes at 4°C,
discarding the top layer as you go.
7.
Pour the top aqueous layer into a new sterile microcentrifuge tube.
8.
Discard the bottom layer.
Step 2: Precipitation of Protein and Cell Debris
1.
Fill a sterile 1.5 ml micro-centrifuge tube with 0.1 volume of
sodium acetate (pH 5.2).
2.
Close the tube and invert it to gently stir the contents.
3.
Incubate the mixture for 15 minutes at -20°C.
4.
Centrifuge the mixture at high speed for 20 minutes at 4°C,
removing the top layer.
5.
Transfer the top aqueous layer into a new sterile microcentrifuge tube.
6.
Discard the bottom layer.
42
Step 3: Precipitation of Nucleic Acids
1.
Fill a sterile 1.5 ml micro-centrifuge tube with 2 volumes of
ice-cold 98% ethanol.
2.
Close the tube and invert it to gently stir the contents.
3.
Incubate the mixture for 15 minutes at -20°C.
4.
Centrifuge the mixture at maximum speed for 20 minutes at
4°C and discard the supernatant.
5.
Add 1 ml of ice-cold 98% ethanol and vortex the mixture
briefly.
6.
Centrifuge the mixture at maximum speed for five minutes
at room temperature and discard the supernatant.
7.
Add 1 cc of ice-cold 70% ethanol and vortex the mixture
8.
Centrifuge the mixture at maximum speed for five minutes
briefly.
at room temperature and discard the supernatant.
9.
Repeat steps 7-10 with 1 ml of ice-cold 70% ethanol
(optional).
10.
Use air to dry the pellet.
11.
Add 10 μl of 1XTE (Option 1) or 10 μl of water (Option 2).
12.
Resuspend the pellet by shaking or vortexing.
The advantage of this method includes the ability to use it in the
presence of substances that break down proteins, such as SDS and
urea, as well as with agents that bind to metal ions, like EDTA, and
reagents that interact with sulfhydryl groups and enzymes like
trypsin or chymotrypsin.
43
Chelex Method
The Chelex method is a process for preparing DNA for PCR, which is
a commonly used method for amplifying DNA. This technique uses a
special resin, called Chelex, to protect the DNA from being damaged
by enzymes called DNAases. These enzymes can break apart DNA,
making it impossible to use for PCR. The Chelex resin binds to
essential elements for DNAases, like magnesium ions, and disables
them, thus safeguarding the DNA from harm.
Stock
Chelex 300 μL
Heating Block
ddH2O
Sterile Forceps
Vortex
Centrifuge tubes
Procedure
The method for using Chelex to extract DNA is as follows:
1.
Obtain 1 spare tube for a control and 3 pre-made tubes
filled with 300 L 10% Chelex from the refrigerator. Handle
the container while wearing gloves, and shake the desired
number of tubes into your gloved hand.
2.
Label each Chelex tube with the sample number it belongs
3.
Activate the heating block and preset it at 95 °C. Water is
to and the date and initials.
poured into the openings.
44
4.
Sterilize forceps by dipping them in ethanol and then waving
them across an alcohol burner's flame to ignite. Repeat this
process twice more.
5.
Remove a small piece of tissue from the sample using sterile
forceps. Place the tissue in the Chelex tube with the correct
label, and screw the cap back on.
6.
Repeat step 5 for each sample, sterilizing the forceps three
times between samples. Create a negative Chelex control by
dipping sterilized forceps into a tube of Chelex slurry.
7.
Vortex the samples in the Chelex slurry for 10 to 15 seconds.
8.
Spin the samples rapidly in a microcentrifuge for 10-15
seconds.
9.
Heat the samples to 95 °C for 20 minutes. While the tubes
are incubating, check to make sure the covers have not
fallen off.
10.
Vortex the samples once again for 10 to 15 seconds.
11.
Spin the tubes rapidly once more to make sure all the
contents are at the bottom of the microcentrifuge tube.
12.
The samples are now usable for PCR.
Advantages and Disadvantages
The Chelex method has benefits such as being cost-effective, fast,
and safe as it does not involve any dangerous chemicals. However, it
has some drawbacks like not working well with blood samples,
producing DNA samples of poor quality, and not being suitable for
DNA analysis using restriction fragment length polymorphism.
45
Differential Method
The differential extraction technique is a method used to separate
sperm cells from other cell types for DNA extraction. This technique,
also known as differential lysis, is commonly used in sexual assault
cases to compare the DNA profiles of the perpetrator and the victim.
The sperm cells are more challenging to lyse compared to other cells
due to the presence of protein disulfide links in their outer
membrane. The differential method includes several steps, including
an optional wash phase, non-sperm cell lysis, and sperm cell lysis.
Principle
Differential extraction, also known as differential lysis, is a method
used to separate DNA from two different types of cells without
blending
their
contents.
It
is
commonly
used
in
forensic
investigations of sexual assault cases, where the DNA from sperm
cells and vaginal epithelial cells is extracted to compare the DNA
profiles of the attacker and the victim. The reason why sperm cells
can survive the extraction process better than epithelial cells is
because of the presence of protein disulfide links in the outer
membrane of sperm cells. These links protect the DNA from being
damaged during the extraction process.
Procedure
Differential Extraction can be carried out as follows:
1.
This is an optional step to remove pollutants and cellular
waste. To carry out this step, add a buffer and detergent to
the sample and incubate it either at room temperature or in
46
a refrigerator. Discard the supernatant (wash fraction) after
the sample has been cleaned.
2.
Mix the sample with an extraction buffer that contains a
buffer, detergent, and proteinase K. Incubate the mixture to
lyse all cells but for spermatozoa. Remove the supernatant
(fraction 1) containing the DNA from the lysed cells and
wash the sperm pellet with a buffer several times to remove
extra DNA.
3.
Incubate the pelleted sperm cells with a buffer, detergent,
DTT, and a stronger dose of proteinase K to lyse the sperm
cells. This will result in fraction 2.
4.
Extract each fraction, including the wash fraction if
necessary, with a phenol/chloroform/isoamyl alcohol mixture
to purify the DNA.
Advantage and Disadvantage
Differential extraction is a method used in forensic science to
separate the DNA from multiple contributors in a sample, such as a
male culprit and a female victim. This process is important because it
helps eliminate confusion and improve accuracy when identifying
the sources of DNA. By following quality assurance standards, the
process ensures the quality of the DNA extraction. However, there is
a potential drawback to differential extraction as the sperm head
must be strong enough to withstand the extraction process or the
results may be unreliable.
47
Qualification of Nucleic Acids
Qualification of nucleic acids refers to the process of characterizing
and verifying the purity and integrity of nucleic acid samples, such as
DNA or RNA. It is an important step in many molecular biology and
biotechnology applications, such as cloning, sequencing, and gene
expression analysis.
There are several methods that can be used to qualify nucleic acids,
including:
Spectrophotometry
This method uses a spectrophotometer to measure the absorbance
of nucleic acids at different wavelengths. The ratio of absorbance at
260 nm to 280 nm (A260/A280) is used to calculate the purity of the
nucleic acids. A ratio of around 1.8 is considered to indicate a pure
sample of DNA, while a ratio of around 2.0 is considered to indicate
a pure sample of RNA.
Gel electrophoresis
This method uses agarose or polyacrylamide gel electrophoresis to
separate nucleic acids based on size and charge. The separated
nucleic acids can then be visualized under UV light after staining with
ethidium bromide or a similar dye. This method can be used to
confirm the size and integrity of the nucleic acids, as well as to detect
contaminants such as proteins or salts.
48
Fluorometry
Fluorometry uses a fluorometer to detect nucleic acids by measuring
their fluorescence. It is a very sensitive method, which can quantitate
the amount of nucleic acid present in a sample. It can also be used
to determine the ratio of double-stranded to single-stranded nucleic
acids, providing information about the integrity and quality of the
sample.
Quantitative PCR (qPCR)
QPCR is a method that uses fluorescent dyes and PCR amplification
to quantify nucleic acids. It can be used to measure the copy number
of specific sequences in a sample, and also to determine the quality
and integrity of the nucleic acids.
Other Methods
Other methods like gel filtration, centrifugation, electrophoretic
mobility shift assay (EMSA) and others can also be used to qualify
nucleic acids.
It's important to note that, depending on the downstream
application, different methods might be more appropriate for
different samples. Therefore, it's important to carefully consider the
sample type and downstream application when choosing which
method(s) to use for nucleic acid qualification.
49
Spectrophotometry
To measure the amount of DNA or RNA in a sample, scientists use a
process called quantitation of nucleic acids. This is important
because reactions that use these types of molecules require a
specific amount and purity for best results. There are different
methods to determine the concentration of nucleic acids, including
spectrophotometry and UV fluorescence.
Spectrophotometry works by exposing the sample to ultraviolet light
and measuring the amount of light that passes through the sample.
The sample absorbs the ultraviolet light in a specific pattern, and the
more light that is absorbed, the higher the concentration of nucleic
acids in the sample. This measurement is done using a machine
called a spectrophotometer and it detects the light absorbed at a
specific wavelength of 260 nm.
Principle
Using the Beer-Lambert Law, it is possible to relate the amount of
light absorbed to the concentration of the absorbing molecule. The
extinction coefficient, which is the amount of light absorbed, is
different for different types of molecules. For example, doublestranded DNA has an extinction coefficient of 0.020 (μg/ml) −1cm−1
at a wavelength of 260 nm, while single-stranded RNA has an
extinction coefficient of 0.025 (μg/ml) −1 cm−1. This means that the
concentration of the absorbing molecule can be calculated based on
the amount of light absorbed. A value of 1 Absorbance (A)
corresponds to a concentration of 50 μg/ml for double-stranded
DNA. However, this method is only valid for up to an A of 2. For
50
more accuracy, a prediction of the extinction coefficient can be made
using the nearest-neighbor model, especially for short singlestranded oligonucleotides which are dependent on the length and
base composition.
Stock
UV/VIS Spectrophotometer
1 ml quartz cuvette
DNA sample(s)
TE Buffer
Disposable 1 ml polyethylene Transfer Pipettes(Berol) [2 per
group]
Eppendorf tubes (1.5 ml) [2 per group]
Ruler
Kim wipes
Procedure
Step 1: Setting up the Spectrophotometer (BeckmanDU64)
1.
Turn on the spectrophotometer at the power strip and
ensure that the printer is connected and ready.
2.
Turn on the UV lamp source and allow it to warm up for 5
minutes.
3.
Select the absorbance reading mode (ABS key). Press the
SCAN key, "Edit" will be displayed.
4.
Enter the starting wavelength as 280 nanometers (nm) and
press enter. Enter the ending wavelength as 260 nm and
press enter.
The speed for the scan of the sample will be displayed. It
should read 750 nm/min. If it does not, press the STEP key
and scroll through the options until 750 nm/min is
displayed. Press enter.
51
5.
Upper limit will be displayed. Set the upper limit at 2,000
absorbance. Press enter.
6.
Lower limit will be displayed. Set the lower limit at 0.000
absorbance. Press enter. The starting wavelength will
reappear.
The instrument is now ready to be calibrated against a
control solution. The purpose of the calibration is to
measure and then subtract from the sample absorbance any
absorbance from the buffer solution.
7.
Place 200 microliters (μL) of the TE Buffer into the quartz
cuvette. This is the solution you will use to calibrate the
instrument.
8.
Open the sample compartment lid on the instrument.
Carefully wipe the cuvette with a Kimwipe and be careful not
to get fingerprints on the quartz panels. Place the cuvette
into the cuvette holder so that the quartz sides are in the
path of the light source (left to right).
9.
Close the sample compartment lid. Press CALB. The
absorbance of the TE Buffer solution will now be recorded in
memory as the "background" and "Bkg" will be displayed.
10.
Press READ. Calibration is complete when "Scan" is
displayed. The instrument is now ready to measure DNA
samples.
11.
Open the sample compartment lid. Discard 200 microliters
of TE Buffer from the cuvette by pouring it out and
disposing of it properly.
12.
Thoroughly rinse the cuvette twice with the TE Buffer
solution and then carefully drain the cuvette onto a Kimwipe
to remove any remaining droplets.
52
Step 2: Sample Preparation
1.
Carefully add 200 microliters of the DNA sample to the
cuvette and place the cuvette in the sample holder of the
spectrophotometer.
2.
Initiate the measurement process by pressing the READ
button. The spectrophotometer will measure and record the
absorbance of the sample between 260 and 280 nanometers
and will then plot the results as a graph on the printer.
3.
Repeat steps 2 and 3 for any other DNA samples that you
have been assigned to analyze.
Advantages
1.
It is easy to perform.
2.
It is cost-effective.
3.
It provides reliable results if done correctly.
Disadvantages
1.
It cannot reliably assess protein contamination.
2.
It does not contribute much error to DNA quantity
estimation.
3.
This method requires a spectrophotometer and cuvettes.
53
Characterization of DNA by
Spectrophotometric Assay and
Melting Temperature (Tm)
To make sure that the extracted DNA samples are suitable for further
experiments, it's important to check their quantity and quality. This is
called characterization and can be done using different methods. In
this experiment, two methods will be used: spectrophotometry and
DNA melting temperature analysis.
Spectrophotometry is a way to figure out how much and how pure
the DNA is. The DNA absorbs light at 260 nm and 234 nm in
ultraviolet light, and the 260 nm is the most important for measuring
the amount of DNA. The ratio of the light absorbed at 260 nm and
280 nm can tell us if there's any protein contamination, since
proteins absorb more light at 280 nm. The ratio of light absorbed at
260 nm and 230 nm can also help us make sure that the sample is
pure and free from other contaminants like carbohydrates, peptides,
and ethanol.
DNA melting temperature analysis is used to see how stable the
DNA is and what its GC content is. This method involves heating up a
diluted DNA solution and measuring the light absorbed at 260 nm as
the two strands of the double helix slowly separate.
1.
To determine the concentration and purity of extracted DNA
using UV spectrophotometer.
54
2.
To determine the DNA melting temperature and GC content
percentage.
Principle
The melting temperature (Tm) is a crucial characteristic of DNA that
determines its stability. It is defined as the temperature at which half
of the DNA molecules are no longer paired. The Tm value is
determined by the length and the GC content of the DNA, which is
the proportion of guanine and cytosine nucleotides. The GC content
is significant for the stability of the DNA and can be measured using
a Tm profile.
To ensure that the extracted DNA samples are suitable for further
use, spectrophotometry and DNA melting temperature analysis are
performed.
These
techniques
are
used
to
determine
the
concentration, purity, and stability of the DNA, ensuring that it meets
the requirements for downstream applications.
Stock
The extracted DNA
0.1 X SSC buffer
Preparation of 20X SSC buffer
Dissolve 175.3 g of NaCl, 88.2 g of sodium citrate dehydrate in 800
ml distilled water. Adjust pH to 7.0 with diluted HCl. Make up the
final volume to 1 L with distilled water.
Procedure
1.
Dilute the extracted DNA in 0.1 X SSC buffer. To do this, take
1 ml of the stock DNA and mix it with 10 ml of the buffer,
resulting in a 1:10 ratio of DNA to buffer.
55
2.
Measure the concentration and purity of the DNA sample
using a spectrophotometer. Place the sample in a quartz
cuvette, and use a second cuvette filled with distilled water
as a blank. Set the spectrophotometer to measure the
absorbance of nucleic acids at a wavelength of 10 mm. The
results will be given in μg/ml.
3.
Alternatively, you can measure the absorbance at three
specific wavelengths (230, 260, and 280 nm) to determine
the concentration and purity of the DNA.
4.
Determine the melting temperature of the DNA. Dilute the
stock DNA to a concentration of 10 μg/ml in 0.1 X SSC
buffer and place it in a quartz cuvette. Fill a separate test
tube with 1 ml of distilled water as a blank. Cover both tubes
with aluminum foil and place them in a water bath at 25°C
for 4 minutes to equilibrate.
5.
Transfer the sample and blank to quartz cuvettes and place
them back in the water bath for 1 minute to equilibrate.
Read and record the absorbance at 260 nm.
6.
Increase the temperature of the water bath to 50°C, 60°C,
70°C, and boiling, and repeat the steps of equilibrating the
sample and blank and reading the absorbance at 260 nm.
Observe the change in absorbance to determine the melting
temperature of the DNA.
Wavelength (nm)
230
Absorbance of DNA
R
P
a
l
260
280
56
Results
A.
Characterization of DNA by Spectrophotometric Assay
(concentration and purity):
Find out the concentration of the DNA samples using the
following equation:
Concentration of DNA (μg/ml) = (A260 / ε L)× Dilution factor
(DF)
Determine the purity of the DNA samples by calculating
A260/A280 and A260/A230 ratios.
B.
Melting Temperature
Temperature (°C )
25
DNA Absorbance at 260 nm
R
P
a
l
50
60
70
Boiling
Plot the value of absorbance vs. temperature and calculate
the Tm for sample DNA.
Find out the GC content of your sample using the following
formula: (G + C)% = (Tm - 69.3) x 2.44.
57
Alternative Method for Characterization
Step 1: Extraction of DNA Sample
1.
Obtain the sample to be tested, such as blood, saliva, or
tissue.
2.
Follow the appropriate DNA extraction protocol to isolate
the DNA.
Step 2: Concentration and Purity Analysis
1.
Transfer a known volume of the extracted DNA solution to a
cuvette.
2.
Load the cuvette into a UV spectrophotometer and run the
analysis.
3.
The spectrophotometer will measure the absorbance of the
DNA sample at 260 nm, which is proportional to the
concentration of DNA.
4.
The ratio of the absorbance at 260 nm to 280 nm
(A260/A280) is used to determine the purity of the DNA. A
ratio of 1.8 or higher is considered pure.
Step 3: Determination of DNA Melting Temperature
1.
Load the DNA sample into a thermocycler and run a melting
temperature (Tm) analysis.
2.
The Tm is the temperature at which half of the DNA is
melted. It can be used as an indicator of DNA quality and
purity.
Step 4: Analysis of GC Content Percentage
1.
Transfer a known volume of the extracted DNA sample to a
2.
Add reagents to the PCR tube, including Taq polymerase
PCR tube.
and primers.
58
3.
Run a PCR reaction to amplify the DNA.
4.
Load the amplified DNA product into an electrophoresis gel
and run the analysis.
5.
The GC content of the DNA can be estimated based on the
position of the DNA fragments on the gel.
Note: The steps outlined above are general and may vary depending
on the specific protocols used in the laboratory.
59
Fast Technology Analysis
Fast technology analysis method is not a specific method that is
used in DNA extraction or purification. It could refer to any
technology or method that allows for faster analysis or processing of
DNA samples.
For example, it could refer to the use of high-throughput sequencing
technologies such as Illumina or PacBio, which can rapidly generate
large amounts of DNA sequencing data. Or it could refer to
automation of lab processes such as using robotic liquid handlers for
DNA extraction and PCR set-up.
In general, fast technology analysis method can be used in various
stages of the DNA analysis process, such as sample preparation,
DNA isolation, amplification, sequencing, and data analysis. The use
of these methods can increase the speed and efficiency of DNA
analysis, resulting in faster and more accurate results.
It's important to note that, while fast technology analysis method
can be very powerful, it's also important to consider the possibility of
errors and artifacts that may occur with these technologies.
Therefore, quality control measures and proper validation are
essential to ensure the accuracy and reliability of the results
obtained.
Paper extraction
The fast technology analysis (FTA) paper extraction method is a
simple technique used to extract DNA, initially in forensic science,
but now widely used in other fields as well. The process involves
60
smearing a sample, typically blood, onto a piece of paper, allowing it
to dry. Then, circles are punched out from the dried paper, and put
into a test tube. The extracted DNA is then cleaned using a solvent,
before finally being added to a polymerase chain reaction (PCR)
mixture. This method is known for its ease and simplicity, which is
why it has gained popularity in various fields.
Principle
The FTA (fluid transfer assay) method is a process used to preserve
and protect DNA in biological samples such as blood and saliva. The
idea behind this method is that when these samples are applied to a
special type of paper, the biological material will stick to the paper
while the chemical mixture used in the process will break down the
cells and change the proteins.
Once the sample has been dried and stored properly, the nucleases,
which are enzymes that break down DNA, are no longer active. This
leaves the DNA stable and protected from damage caused by factors
such as oxidation, ultraviolet light, bacteria, and fungus. The FTA
method helps to minimize the damage to nucleic acids and ensures
that the DNA remains intact for further analysis.
Stock
FTA Purification Reagent
TE Buffer (10mM Tris-HCL, 0.1 mM EDTA, pH 8.0)
Solution 1:0.1N NaOH, 0.3mM EDTA, pH13.0
Solution 2:0.1M Tris-HCL, pH 7.0
Procedure
Step 1: Washing
61
1.
Obtain a 6 mm punch from a dry area using a standard
single-hole paper punch.
2.
Place the punch in a 1.5 ml microtube and rinse the cutting
end with ethanol to prevent cross-contamination.
3.
Pour 1000 μl of the FTA purification reagent into the
microtube and mix the mixture by either manual stirring or
flash vortexing five times.
4.
Allow the mixture to sit at room temperature for 5 minutes,
and mix it again if desired.
5.
Using a pipette, remove and discard all used FTA purification
reagent.
6.
7.
Repeat steps 2-4 a total of three times.
Add 1000 μl of TE buffer to the microtube and incubate at
room temperature for 5 minutes.
8.
Using a pipette, remove and discard all used TE Buffer.
9.
Repeat steps 6-8 twice more for a total of three washes with
TE buffer. The punch should be white or pale in color after
this process.
Step 2: pH Treatment
1.
Obtain a 6 mm punch that has been previously rinsed and
add 140 μl of Solution 1.
2.
Incubate the punch at 65°C for 5 minutes (this is different
from the typical room temperature protocol used by the FTA
company).
3.
Add 260 μl of Solution 2 and mix the mixture by flashing the
4.
Allow the mixture to sit at room temperature for an
vortex five times.
additional 10 minutes.
5.
Vortex the mixture again for 10 flashes.
6.
Remove the punch and squeeze it to recover the most elute
volume possible.
62
7.
Use a clean pipette tip to remove the punch if needed.
8.
The elute carries 66 mM Tris-HCl, 0.1 mM EDTA. For a 25 μl
PCR reaction, use 0.5 μl of the elute.
Advantage
The FTA method has several benefits that make it an attractive
option for storing and preserving DNA. Firstly, it does not require
refrigeration and can be kept at room temperature, making it
convenient for storage and transportation. Secondly, it is effective in
killing harmful bacteria while preserving the DNA, ensuring its quality
and integrity. The small size of the discs makes it easy to store and
ship, and the process of obtaining the DNA is simple and can be
repeated multiple times without having to measure the amount of
DNA beforehand.
Disadvantage
However, there is also one disadvantage to the FTA method - the
small discs of DNA can be easily contaminated by static electricity.
This means that there is a risk of contamination during handling,
which could impact the quality of the DNA obtained using this
method.
63
Recombinant DNA Technology
Recombinant DNA technology, also known as genetic engineering, is
the process of manipulating the genetic makeup of an organism by
inserting, deleting or replacing specific genes. This technology allows
for the production of genetically modified organisms (GMOs) with
desired traits, such as resistance to disease or improved nutritional
content.
The technology comprises the extraction of a specific gene or DNA
segment from an organism, which is then inserted into a vector like a
plasmid. The vector is then introduced into a host organism, which
can be a plant, animal, yeast, or bacteria. Once inside the host
organism, the vector replicates and expresses the inserted DNA.
This technology has numerous applications across various fields such
as agriculture, medicine, and biotechnology. For instance, in
agriculture, GMOs with pest and disease resistance can be produced,
leading to increased crop yields. In medicine, it is utilized to produce
human
insulin
and
other
therapeutic
proteins,
while
in
biotechnology, it is used for enzyme and industrial product
production.
However, the use of recombinant DNA technology also raises safety
and ethical concerns, such as potential unintended effects on the
environment and human health from the modification of food crops.
There are also ethical concerns regarding the use of recombinant
DNA technology in medical treatments, like gene therapy. Hence, it's
crucial that any application of recombinant DNA technology
64
undergoes thorough evaluation for safety and ethical implications
and is continuously monitored and regulated for the future.
65
Designing DNA Probes
DNA probe designing is the process of creating a DNA fragment that
can selectively hybridize to a complementary DNA target sequence.
DNA probes are used to detect and identify specific DNA sequences
in a sample. The design of a DNA probe involves selecting a
sequence of DNA that is complementary to the target sequence, as
well as optimizing the length, specificity, and sensitivity of the probe.
Different types of probes can be designed, including fluorescent,
biotinylated, or radioactive probes, depending on the intended use.
DNA probe designing is an essential step in many molecular biology
techniques, such as polymerase chain reaction (PCR), DNA
sequencing, and gene expression analysis.
Principle
The principle of DNA probe design is to create a short, singlestranded piece of DNA that is complementary to a specific sequence
of the target DNA. This complementary sequence allows the probe
to hybridize or bind with the target DNA, allowing for detection or
identification of the target sequence. The design of the DNA probe
must also consider factors such as length, specificity, and sensitivity
to ensure accurate and reliable results.
Procedure
1.
Determine the target DNA sequence by searching databases
or sequencing DNA of interest.
2.
Decide on the length of the DNA probe, aiming for 18-25
nucleotides. The length of the DNA probe is an important
66
consideration as it can affect the specificity and sensitivity of
the probe.
3.
Choose a labeling method such as fluorescent dyes, biotin,
or radioactive isotopes. The choice of labeling method will
depend on the intended use of the probe and the detection
method being used.
4.
Design the probe sequence to complement the target DNA
sequence and include the chosen labeling moiety. The probe
sequence should be complementary to the target DNA
sequence and should contain the labeling moiety at the
appropriate location. It is important to avoid secondary
structures and self-complementarity in the probe sequence.
5.
Test the probe for specificity and sensitivity by hybridizing it
to the target DNA sequence.
6.
Optimize the probe by adjusting its length, labeling method,
or concentration.
7.
Validate the probe by testing it on a variety of samples and
verifying consistent and reproducible results.
Advantages
1.
DNA probes are designed to be complementary to a specific
DNA sequence, so they are highly specific in their detection
of the target DNA.
2.
DNA probes can detect even small amounts of the target
DNA, making them a powerful tool in molecular biology.
3.
DNA probes can provide rapid results, with detection often
4.
DNA probes can be designed to detect any DNA sequence,
taking only a few hours.
making them a versatile tool for a range of applications.
67
Disadvantages
1.
DNA probes can be expensive to design and manufacture,
particularly if multiple probes are required for a single
experiment.
2.
Designing a DNA probe requires a high level of expertise in
molecular biology, and the process can be complex and
time-consuming.
3.
DNA probes can sometimes cross-react with other DNA
sequences, leading to false positive results.
4.
DNA probes can degrade over time, so they must be stored
carefully and used quickly to ensure accuracy.
68
Bergs Terminal Transferase - Boyer
Cohen Chang Experiment
Berg's Terminal Transferase (TdT) is an enzyme that catalyzes the
addition of deoxy-nucleotides to the 3' end of DNA strands. The
Boyer-Cohen-Chang
experiment
was
conducted
in
1973
to
demonstrate the activity of TdT. The experiment involved incubating
a
partially
double-stranded
DNA
template
with
TdT,
deoxynucleotides, and radioactive ATP. The results showed that TdT
added radioactive nucleotides to the 3' end of the DNA strand,
proving its ability to extend DNA chains. This experiment was critical
in establishing TdT's role in DNA synthesis and repair processes.
Principle
The principle of Berg's Terminal Transferase – Boyer Cohen Chang
experiment is to use a bacterial enzyme called terminal transferase to
add homopolymeric tails to the 3' ends of DNA molecules. This
process is important for many molecular biology techniques, such as
cloning and sequencing, as it allows for specific and efficient
hybridization of DNA molecules. The experiment demonstrated that
the terminal transferase enzyme could add homopolymeric tails to
double-stranded DNA, and that the length and composition of the
tails could be controlled by adjusting the reaction conditions. The
discovery of this enzyme and its ability to modify DNA has greatly
facilitated the development of many molecular biology techniques.
69
Procedure
1.
Prepare a reaction mixture containing the following
components - DNA template, terminal deoxynucleotidyl
transferase (TdT), dGTP, dATP, dTTP, and MnCl2.
2.
Mix the reaction mixture and incubate it at 37°C for 15-60
minutes.
3.
Stop the reaction by adding EDTA.
4.
Denature the DNA by heating the reaction mixture at 95°C
for 5 minutes.
5.
Analyze the reaction products using polyacrylamide gel
electrophoresis.
6.
Visualize the reaction products by staining the gel with
7.
Compare the size and intensity of the reaction products with
ethidium bromide.
the control samples.
8.
Interpret the results and draw conclusions based on the
findings.
9.
Repeat the experiment with different variations to confirm
the results.
Advantages
1.
Berg's terminal transferase (TdT) is a highly specific enzyme
that can incorporate nucleotides onto the 3' end of DNA
fragments.
2.
The Boyer Cohen Chang experiment provided a new and
efficient method for introducing DNA into bacterial cells,
which revolutionized the field of genetic engineering.
3.
TdT has low error rates, making it an ideal tool for
constructing cDNA libraries.
70
4.
The Boyer Cohen Chang experiment paved the way for
recombinant DNA technology, which has since been used to
produce a wide range of useful products.
Disadvantages
1.
TdT is highly sensitive to temperature, pH, and salt
concentration, which can lead to variable results and limit its
use in certain applications.
2.
The Boyer Cohen Chang method requires the use of
antibiotics to select for transformed cells, which can lead to
the development of antibiotic-resistant strains.
3.
The process of creating recombinant DNA can be timeconsuming and complex, requiring specialized equipment
and expertise.
4.
There are ethical
concerns
surrounding the use of
recombinant DNA technology, particularly with regards to
genetically modified organisms (GMOs).
71
Preparation of Competent Cells for
Efficient DNA Uptake in E. coli
Competent cells are a type of laboratory-grown bacterial cells that
have been artificially modified to accept foreign DNA into their
genomes. The process of modifying the cells to be competent
involves making small changes to their cell membranes and cellular
components, which allows them to efficiently uptake and integrate
foreign DNA.
Competent cells are an essential tool in molecular biology,
biotechnology, and genetic engineering. They are used in a variety of
applications including gene cloning, genetic modification, and the
production of recombinant proteins. The ease and efficiency of DNA
transfer into competent cells makes them a crucial tool in the field of
biotechnology and genetic engineering.
One of the most common methods for making competent cells is by
transforming them with calcium chloride. In this process, the bacteria
are treated with a high concentration of calcium chloride, which
temporarily permeates the cell membrane, allowing foreign DNA to
enter. Once the DNA has been taken up by the cells, they return to
their normal state, and the DNA becomes integrated into the
genome.
Another method for making competent cells is by electroporation,
which involves applying a high-voltage electrical pulse to the cells,
which creates temporary pores in the cell membrane, allowing the
foreign DNA to enter. This method is often faster and more efficient
72
than the calcium chloride method, but it also involves a higher risk of
cell death.
Competent cells are also classified into two types: chemically
competent cells and electro-competent cells. Chemically competent
cells are made by using a chemical agent, such as calcium chloride,
to permeate the cell membrane and allow DNA to enter. Electrocompetent cells are made by using an electrical pulse to create
temporary pores in the cell membrane, allowing the DNA to enter.
In addition to their use in biotechnology and genetic engineering,
competent cells also have applications in medical research. They can
be used to produce vaccines, produce recombinant proteins for use
in diagnostic tests, and to study the interactions between bacteria
and their hosts.
Procedure
1.
Choose a bacteria that is susceptible to transformation, such
as E. coli, and grow it in a culture medium until it reaches
logarithmic phase.
2.
Centrifuge the bacterial culture to harvest the cells. Discard
the supernatant and resuspend the pellet in cold sterile
buffer.
3.
Slowly freeze the cells in a solution of glycerol and buffer.
This will protect the cells from damage and make them more
competent for transformation.
4.
Thaw the cells quickly in a water bath at 37°C. The cells
should be kept at 37°C for no more than 30 seconds.
5.
Add calcium chloride to the cells to increase their
competence. Incubate the cells at 37°C for 15 minutes.
73
6.
Add the DNA to be transformed to the competent cells.
Incubate the cells at 37°C for 1 hour.
7.
Transfer the transformed cells to a nutrient agar plate and
incubate overnight at 37°C.
8.
Screen the colonies that have grown on the agar plate for
the desired phenotype. Confirm the transformation by
conducting further tests, such as PCR or sequencing.
Alternative Method
Principle
Competent cells in bacteria are necessary for efficient DNA uptake.
The bacteria's plasma membrane must be permeable to foreign
DNA, which can be achieved by artificially making the bacterial cells
competent. However, since both bacteria and DNA are negatively
charged, it is challenging for DNA to enter bacterial cells. To
overcome this, divalent cations are used, which shield charges by
coordinating phosphate groups and other negative charges, thereby
changing the electrical charge on the bacterial cell surface and
weakening the bacterial cell membrane. This promotes the entry of
foreign DNA into bacterial cells.
Stock
Bacteria
Spectrophotometer
LB Agar Plate
LB Broth
Glycerol
CaCl2
Tubes
Centrifuge
74
Preparation of LB Medium
Prepare Luria Bertani medium by mixing 10 g tryptone, 5 g yeast
extract, and 10 g of NaCl in 1 litre of distilled water. Adjust the pH to
7.0 with 1N NaOH and autoclave the mixture for 25 minutes at
120°C.
Procedure
1.
Grow E. coli in 5 ml LB Broth overnight.
2.
Dilute overnight culture 100-fold and grow bacteria in 250
ml of LB in 1L conical flask at 37°C, over a shaker.
3.
Check the OD of the culture at 600 nm until it reaches 0.250.30.
4.
Transfer culture into 50 ml centrifuge tubes and centrifuge at
5.
Discard supernatant and resuspend cells in 30 ml of chilled
3,000 rpm for 10 minutes at 4°C.
100 mM CaCl2 for 30 minutes on ice with intermittent hand
shaking.
6.
Centrifuge cells at 3,000 rpm for 10 minutes at 4°C, discard
supernatant, and resuspend bacterial pellet in 5-10 ml of 100
mM CaCl2 plus 15% glycerol.
7.
Prepare 100 ul aliquots of this bacterial cell suspension and
store at -70°C for further use.
Observation
Competent cells are prepared successfully and are frozen at -70°C
for further use.
75
Precautions
1.
Perform all procedures in cold conditions.
2.
After CaCl2 treatment, bacterial cells become fragile, so avoid
vortexing them.
3.
Do not freeze and thaw competent cells repeatedly.
76
Bacterial Transformation in vitro
using Electroporation
Principle
Plasmid DNA is a circular, double-stranded DNA molecule that exists
independently from the chromosomal DNA of bacterial cells.
Plasmids often contain genes that can provide an advantage to the
bacterial cell, such as antibiotic resistance genes. Introducing
plasmid DNA into a bacterial cell provides the cell with new genetic
material that can give it a new function or advantage.
Bacterial transformation is the process of introducing foreign DNA
into bacterial cells, which can be achieved through various methods
such as electroporation, heat shock, and chemical transformation. In
this experiment, electroporation is used to introduce plasmid DNA
into bacterial cells by creating transient pores in the cell membrane
using a high voltage electric pulse.
Stock
Bacterial culture
Plasmid DNA
Restriction enzymes
DNA ligase
LB agar plates
Ampicillin
Sterile pipettes and tips
Microcentrifuge tubes
Heat block
77
Incubator
Electroporator
Procedure
1.
Prepare bacterial cells for electroporation by centrifuging the
culture and resuspending the cells in ice-cold sterile water.
2.
Wash the cells twice to remove any residual media.
3.
In a microcentrifuge tube, mix 50 μl of the washed cells with
2 μl of linearized plasmid DNA.
4.
Transfer the cell and plasmid DNA mixture to a sterile
electroporation cuvette.
5.
Apply an electric pulse to the cells using the electroporator
according to the manufacturer's instructions.
6.
Immediately add 1 ml of LB broth to the cuvette and transfer
the mixture to a sterile microcentrifuge tube.
7.
Incubate the cells at 37°C for 1 hour to allow for expression
of the plasmid genes.
8.
Plate the transformed cells on LB agar plates containing
ampicillin and incubate overnight at 37°C.
9.
Observe bacterial growth and look for colonies that have
grown on the ampicillin-containing plates.
Results
Successful transformation will result in colonies growing on the
ampicillin-containing plates, which are resistant to the antibiotic and
carry the plasmid DNA. The presence of the plasmid DNA can be
confirmed by performing a plasmid isolation and restriction digest,
or by sequencing the plasmid.
78
Precautions
1.
Maintain aseptic techniques throughout the experiment to
prevent contamination of the bacterial culture with other
microorganisms or foreign DNA.
2.
Use sterile equipment and solutions to avoid introducing
unwanted DNA or bacteria.
3.
Ensure the bacterial culture is healthy and in the exponential
growth phase to increase the likelihood of transformation.
4.
Use appropriate controls, including an untransformed
control, to verify that any observed changes in bacterial
phenotype are due to the introduction of plasmid DNA and
not an artifact of the experiment.
5.
Optimize electroporation parameters (such as voltage, time
constant, and pulse duration) to maximize the efficiency of
transformation while minimizing cell damage.
6.
Use the appropriate antibiotic concentration to select for
transformed cells without causing any adverse effects on the
growth of the bacterial culture.
7.
Store and handle the plasmid DNA properly to prevent
degradation or contamination.
79
CaCl2 Mediated Transformation
CaCl2 mediated transformation is a method of introducing foreign
DNA into a host organism, often with the aim of expressing a new
trait or function. This transformation method is widely used in
molecular biology, biotechnology, and genetic engineering for the
study and manipulation of genes and genomes.
The transformation process includes the preparation of the DNA to
be introduced into the host organism. The DNA is first extracted
from the source organism, often using a restriction enzyme to cut
the DNA at specific sites. The DNA is then ligated to a plasmid, a
circular piece of DNA that is used as a vector for transformation. The
plasmid contains a gene of interest and the origin of replication,
which allows the plasmid to replicate in the host organism.
Once the plasmid is ready, the next step is to introduce it into the
host organism. This is where CaCl2 comes into play. CaCl2 is used as a
mediator in the transformation process to help the plasmid enter the
host organism. CaCl2 acts as a permeater, making the cell membrane
of the host organism temporarily permeable, allowing the plasmid to
enter the cell.
The transformation process can be carried out in various ways, such
as electroporation, where an electrical field is applied to the host
organism, or chemical transformation, where chemicals such as CaCl 2
are used to create temporary holes in the cell membrane. In both
methods, the permeated cells are then incubated with the plasmid
solution, allowing the plasmid to enter the cell.
80
Once the plasmid is inside the cell, it can integrate into the host
organism’s genome or remain as a separate piece of DNA. If the
plasmid integrates into the host organism’s genome, the new gene
will be expressed and the host organism will display the new trait or
function. If the plasmid remains as a separate piece of DNA, it can
still express the new gene and the host organism will still display the
new trait or function.
Procedure
1.
Grow bacterial cells in liquid medium until they reach midlog phase, then chill the culture on ice for 10 minutes.
Transfer the cells to 50 ml tubes containing ice-cold 50%
glycerol and mix gently by inverting the tubes. Store the
competent cells at -80°C until use.
2.
Isolate the plasmid DNA and purify it using a plasmid
isolation kit.
3.
Dissolve 0.1 M CaCl2 in distilled water and sterilize the
solution by autoclaving.
4.
Thaw the competent cells on ice and add the purified
plasmid DNA to the cells. Incubate the mixture on ice for 30
minutes.
5.
Add the sterilized CaCl2 solution to the cell-DNA mixture,
keeping the volume of CaCl2 solution equal to that of the
cell suspension. Incubate the mixture on ice for another 2-5
minutes.
6.
Transfer the mixture to a 42°C water bath and incubate it for
exactly 90 seconds. Quickly transfer the mixture back to the
ice and incubate it for 2 minutes.
81
7.
Add 1 ml of LB medium to the mixture and incubate it at
37°C for 1 hour. Spread the mixture onto an LB agar plate
containing the appropriate antibiotics and incubate it
overnight at 37°C.
8.
Incubate the plate for an additional 16-20 hours at 37°C to
allow for the growth of the transformed cells. Check for the
presence of colonies and pick the desired colonies for
further analysis.
82
Agarose Gel Electrophoresis
Agarose gel electrophoresis is a laboratory technique used to
separate and analyze different sizes of DNA fragments. The process
includes the preparation of a gel from agarose, a sugar found in
seaweed. This gel is poured into an electrophoresis chamber and
solidified. Then, the DNA samples are loaded into wells in the gel. An
electric field is then applied to the mixture, causing the negatively
charged DNA molecules to move towards the positively charged
anode. The speed at which the DNA moves through the gel is
dependent on the size of the DNA molecules, allowing for separation
and quantification of different sized fragments.
The DNA bands can be visualized by staining the gel with a
fluorescent dye – ethidium bromide, and observing it under
ultraviolet light. By using a transluminator and a UV light source with
a wavelength of 254 nm, 310 nm, or 354 nm, electrophoresis enables
the separation and localization of DNA in agarose gel. Various
buffers, such as TAE and TBE, can be used in the electrophoresis
process, with different buffers affecting the migration rate of the
DNA. The molecular weight of the DNA, the concentration of
agarose, the conformation of the DNA, and the applied current all
have an impact on the electrophoretic mobility of DNA through
agarose gel.
The size of the DNA fragments that can be separated depends on
the concentration of agarose in the gel, with different concentrations
allowing for the separation of different sized fragments. For example,
a 0.5% gel can separate DNA fragments between 1 kb and 30 kb,
83
while a 2.0% gel can separate fragments between 50 bp and 2 kb
(refer table below):
w/v % Gel type
Size of DNA fragments (1 Kb = 1000 bp)
0.5 %
1 kb to 30 kb
0.7 %
800 bp to 12 kb
1.0 %
500 bp to 10 kb
1.2 %
400 bp to 07 kb
1.5 %
200 bp to 03 kb
2.0 %
50 bp to 02 kb
Agarose gel electrophoresis is widely used in molecular biology and
genetics and has several applications, including the analysis of PCR
products, the separation of DNA fragments after restriction
digestion, and the detection of genetic variations such as mutations
and polymorphisms. It is important to note that for the separation of
fragments smaller than 100 bp, a different technique called
Polyacrylamide gel electrophoresis is used.
1.
To assess the purity of the extracted DNA by agarose gel
electrophoresis.
2.
To separate and calculate the molecular size of DNA
fragment by comparing the separated bands with known
standard molecular weight marker.
3.
To quantify DNA fragment by comparing the separated
band with known quantity of DNA.
Principle
Electrophoresis is a laboratory method used to separate charged
molecules, like DNA, based on their size and charge. An electric
current is applied to the DNA sample, previously broken down by
restriction enzymes, driving the negatively charged DNA towards the
positive electrode. The agarose gel acts as a semi-solid matrix that
84
allows the DNA fragments to separate based on their size. Larger
molecules travel further, while smaller ones travel less distance. This
enables effective separation of the biomolecules. The separated
fragments can then be analyzed and visualized. The equipment
typically includes a power source, a casting tray for the gel, and a
container for the sample.
Stock
1.
Tris-Borate-EDTA (TBE) stock solution (5X)
Tris base 54.0 gm
Boric acid 27.5 gm
EDTA (pH 8.0) 0.5 M 20.0 ml
Distilled water to make 1000.0 ml
2.
Working Buffer: 1X or 0.5X TBE
3.
Loading Buffer (10X)
Bromophenol blue 0.25%
Xylene cyanol 0.25%
Ficoll (type 400) 25.0%
in distilled water
4.
DNA Sample
DNA 150-200 ng
Distilled water 18 μl
10X loading buffer 2 μl
Before conducting an electrophoresis experiment, it is important to
prepare a stock solution of ethidium bromide. To do this, prepare a
stock solution of 10 mg/ml ethidium bromide and keep it stored at
4oC in a coloured glass tube or container.
85
Agarose powder, 1X TBE buffer (89 mM Tris-base, 89 mM boric acid
and 2 mM EDTA) prepared from 10X TBE, Ethidium Bromide (5
mg/ml), Gel loading dye (Glycerol and orange dye), 1 kb and 100 bp
DNA ladder, horizontal electrophoresis apparatus and power supply.
Procedure
Step 1: Preparation of Agarose Gel
1.
Weigh out the required amount of agarose powder to make
a 1% gel using a scale.
2.
Heat the agarose powder in a microwave to dissolve it and
create a homogeneous mixture.
3.
Add 4 microliters of ethidium bromide to the mixture and
mix it in carefully. This will stain the DNA for visualization
under UV light.
4.
Prepare the gel plate and comb by placing the comb in the
slots on each side of the gel plate and pour the melted
agarose onto the gel plate in the electrophoresis tray.
5.
Let the gel cool to room temperature and remove the comb.
6.
Place the gel in the electrophoresis chamber and add
enough electrophoresis buffer (1X TBE) to cover the gel.
Step 2: Loading of DNA
1.
Mix 300 ng of DNA with 3-4 microliters of loading dye.
2.
Add a DNA ladder (3 microliters) to the first well using a
micro-pipette.
3.
Add the prepared DNA samples to adjacent wells.
4.
Run the electrophoresis at 95 volts for 45 minutes and
5.
Once the electrophoresis is complete, place the gel on a UV
periodically check the gel.
light box to take a picture.
86
Step 3: Observing the Gel
1.
Check the gel under UV light using a transluminator.
2.
Wear a perspex shield or safety glasses to avoid the
damaging effects of UV light.
Note: A lambda DNA digested with Hind III restriction enzyme is
typically used to determine the molecular weight of the experimental
DNA in this procedure. This results in 8 bands of different sizes.
However, similar amounts of degraded genomic DNA will exhibit a
smear instead of sharp bands.
Advantage and Disadvantage
The method of using Ethidium bromide is simple and easy to follow.
It is also known for its fast results and efficient performance.
However, Ethidium bromide is recognized as a carcinogen and thus,
it needs to be used with caution and proper disposal.
87
Protein Analysis by SDS-PAGE
Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis (SDSPAGE), is a widely used method in biochemistry and molecular
biology for the separation and analysis of proteins based on their
size and charge.
The process comprises the denaturation of proteins using SDS, a
detergent that uniformly coats and unfolds the protein, creating a
negative charge proportional to its size. The SDS-protein mixture is
then loaded onto a polyacrylamide gel, which serves as a sieve to
separate the proteins based on their size. An electric field is applied,
causing the negatively charged proteins to migrate towards the
positive electrode, resulting in a separation of the proteins by size.
One of the key advantages of SDS-PAGE is its ability to separate
proteins based on their molecular weight, allowing for easy
identification of specific proteins and the estimation of their relative
molecular weight. This technique is commonly used in the analysis of
protein samples from tissues, cells, and bodily fluids, as well as for
purifying and isolating specific proteins.
SDS-PAGE is also widely used for quality control in biotechnology
and pharmaceutical industries, where it is used to ensure the purity
and consistency of protein-based products such as enzymes,
vaccines, and antibodies. The technique is also used in protein
purification, as well as in the detection of protein modifications such
as phosphorylation, glycosylation, and proteolytic cleavage.
To enhance the resolution of SDS-PAGE, variations such as twodimensional gel electrophoresis (2D-PAGE) can be used. This
88
technique separates proteins based on both size and charge,
providing a more comprehensive view of the protein profile in a
sample.
Stock
Ammonium per sulfate (10%)
Coomassie brilliant blue (0.3%)
Destaining mixture
Gel staining dish
Electrophoresis apparatus with power supply
Running buffer
SDS (10%)
Tris-HCl (1.5 M, pH 8.8)
Tris-HCl (0.5 M, pH 6.8)
Acrylamide-bis-acrylamide stock solution
Reagent Preparation
Acrylamide-bis-acrylamide stock solution: Dissolve 29.2 g
acrylamide and 0.8 g bis-acrylamide in distilled water, then
raise the final volume to 100 ml.
Tris-HCl (1.5 M, pH 8.8): Dissolve 18.15 g of Tris in 50 ml
distilled water, adjust the pH to 8.8 with HCl, then raise the
final volume to 100 ml.
Tris-HCl (0.5 M, pH 6.8): Dissolve 6 g Tris in 60 ml distilled
water, adjust the pH to 6.8 with HCl, then raise the final
volume to 100 ml.
SDS (10%): Dissolve 1 g SDS in 5 ml distilled water, then raise
the final volume to 10 ml.
89
Gel running buffer: Dissolve 14.4 g glycine and 1 g SDS in 1 L
distilled water, adjust the pH to 8.3 with Tris, then raise the
final volume to 1 L.
Ammonium per sulfate (APS) (10%): Dissolve 500 mg solid
APS in 5 ml distilled water. Use freshly prepared APS solution
only.
Coomassie Brilliant Blue R 250: Dissolve 600 mg CBBR-250 in
80 ml methanol, add 20 ml glacial acetic acid, then raise the
final volume to 200 ml with distilled water.
Destaining solution: Mix 400 ml methanol, 100 ml glacial
acetic acid, and 500 ml distilled water to obtain 1 L of
solution.
Laemmli buffer: 62.5 mM Tris-HCl, pH 6.8 (diluted from 0.5
M Tris-HCl, pH 6.8), 10% glycerol, 5% mercaptoethanol, 2%
SDS.
Procedure
SDS-PAGE is a commonly used technique for separation and analysis
of proteins based on their molecular weight. Here is a step-by-step
procedure to perform SDS-PAGE of a protein sample:
1.
To ensure efficient separation, it is important to denature
and reduce the protein sample. This is typically done by
adding a detergent like SDS to the sample and heating it to
95-100°C for 5-10 minutes. In addition, a reducing agent like
DTT or beta-mercaptoethanol can be added to reduce any
disulfide bonds present in the protein sample.
2.
The denatured and reduced protein sample is mixed with a
loading buffer which contains SDS, glycerol, and a tracking
dye to monitor protein migration during electrophoresis.
90
3.
An electrophoresis apparatus, such as a vertical or horizontal
gel apparatus, is set up and the acrylamide gel is prepared
and poured into the apparatus. The gel is then allowed to
polymerize.
4.
Using a micropipette, load the denatured protein solution
into the wells. The protein sample should be denatured in
Laemmli buffer by boiling for 5 minutes. Add standard
molecular weight marker proteins in one lane. For detection
by CBB dye, 20 to 50 g protein is generally sufficient.
5.
Connect the electrodes of the apparatus tightly to the power
supply and run the gel at a constant current of 20 mA.
Larger proteins move slower through the gel compared to
smaller proteins. Track the mobility of the sample in the
matrix using dye (bromophenol blue is generally added to
the Laemmli buffer). The tracking dye provides a visual
representation of the migration of proteins. Once the run is
complete, switch off the button and disconnect the
apparatus.
6.
Transfer the gel to a staining tray containing a protein stain
like Coomassie blue or silver stain to visualize the separated
protein bands. Stain the gel for at least 2 hours or overnight
under shaking conditions on a rocking shaker. The entire gel
should turn blue.
7.
Carefully transfer the gel to a de-staining solution and shake
on a rocker shaker for 30 minutes. Add fresh de-staining
solution, repeating these steps until the bands are clearly
visible in the gel. At this stage, take a photograph of the gel.
8.
Analyze the photographed gel to observe several distinct
blue-colored bands. Each band represents one or multiple
bands in the lane, with the intensity of these bands varying
depending on the amount of polypeptide present in the
protein solution loaded in the gel.
91
Blue-White Colony Selection
employing X-Gal / IPTG
Blue-white colony selection is a widely used method in molecular
biology to select transformants, which are cells that have taken up a
plasmid with a desired gene or sequence. This method is based on
the use of X-Gal, a blue chromogenic substrate, and IPTG, an inducer
of gene expression, to visualize and select colonies that contain the
desired sequence.
The
process
of
blue-white
colony
selection
involves
the
transformation of bacteria with a plasmid that contains a gene of
interest. The plasmid also contains a lacZ gene, which encodes for βgalactosidase - an enzyme that hydrolyzes X-Gal to produce a blue
color. To ensure that the lacZ gene is expressed only when the gene
of interest is present, the plasmid also contains a promoter that is
regulated by IPTG.
After the transformation, bacteria are plated on agar plates that
contain X-Gal and IPTG. Bacteria that have taken up the plasmid will
express the lacZ gene and produce β-galactosidase, which will
hydrolyze X-Gal to produce blue colonies. Bacteria that have not
taken up the plasmid will not express the lacZ gene, and therefore
will not produce β-galactosidase, resulting in white colonies.
The presence of IPTG in the agar medium will induce the expression
of the lacZ gene, leading to the production of β-galactosidase and
the hydrolysis of X-Gal to produce blue colonies. The concentration
of IPTG used in the agar medium can be adjusted to control the level
92
of gene expression and ensure that only the desired colonies are
selected.
The blue-white colony selection method has several advantages over
other selection methods. Firstly, it is simple and easy to perform,
requiring only the addition of X-Gal and IPTG to the agar medium.
Secondly, it is rapid, allowing the selection of colonies within a few
hours. Finally, it is highly specific, as only colonies that contain the
desired gene or sequence will be blue. Its simplicity, rapidity, and
specificity make it an ideal choice for a variety of applications,
including cloning, gene expression, and protein production.
Stock
X-Gal
Dimethylformamide (DMF)
dH2O
Isopropyl β-D-1-thiogalactopyranoside (IPTG)
Screening Antibiotic
Agar Media
Plates.
Procedure
1. Preparation of X-Gal and IPTG:
•
To incorporate X-Gal and IPTG, prepare a 20 mg/ml X-Gal
solution in DMF and a 100 mM IPTG solution in dH2O (see
IPTG Stock Solution Procedure) or dilute from a 1M IPTG
solution.
•
Integrate X-Gal and IPTG into the agar media by adding 10 μl
of 20 mg/ml X-Gal solution per 1 ml of media or 2 μl of 100
mg/ml X-Gal solution per 1 ml of media.
•
To obtain a final concentration of 1 mM IPTG, add 10 μl of 100
mM IPTG solution per 1 ml of media.
93
Notes
To enhance the screening process, a higher concentration of X-Gal
may be used. It increases blue color intensity, reduces blue color
development time, refrigeration time, and decreases the number of
ambiguous colonies requiring rescreening.
2. Screening on agar media containing IPTG and X-Gal:
•
Autoclave the growth media agar and cool to 50°C.
•
Add the screening antibiotic and pour plates. Allow them to
cool to room temperature before use, which usually takes at
least 30 minutes.
Spread transformed competent cells as desired.
•
3. Screening on pre-made agar plates lacking IPTG and X-Gal:
•
Pour autoclaved growth media containing screening antibiotic
on media plates and dry in a laminar flow hood.
•
Add 40 μl of 100 mM IPTG and 120 μl of X-Gal (20 mg/ml) to
the surface of each plate and spread over the entire surface.
•
Dry X-Gal/IPTG-coated media in a laminar flow hood for
approxi-mately 30 minutes before use.
•
Spread transformed competent cells and incubate inverted at
37°C until blue colonies form (usually ~24 hours).
Result
The bacterial cell that is transformed with a vector containing
recombinant DNA will produce white colonies whereas bacterial cell
that is transformed with the vector without recombinant DNA will
produce blue colonies.
94
Notes
In this, foreign DNA is inserted to interrupt the beta-galactosidase
coding sequence, which otherwise reacts with x-gal substrate to
produce a blue color. Due to the defective enzyme produced by
insertion of foreign DNA in gal gene, white colored colonies are
produced.
95
Genomes and Interrelatedness
Genomes refer to the complete set of genetic material that exists
within an organism's cells. It is the blueprint of life, and it carries the
information required for an organism to develop and function. The
genome is made up of DNA, which is a long chain of nucleotides.
These nucleotides store the genetic information in the form of four
letters: A, T, C, and G, which represent the four types of nitrogenous
bases in the DNA molecule.
The concept of genomic interrelatedness is crucial in genetics and
genomics, as it refers to the tight connections between species and
their genomes. This interrelatedness provides insight into the
evolutionary history of life on earth and the relationships between
species.
One of the primary ways that interrelatedness is studied is through
comparative genomics. This involves comparing the genomes of
different species to identify similarities and differences. This
approach provides insight into the evolutionary relationships
between species, including when species evolved and how they are
related. For example, comparing the genomes of humans and
chimpanzees provides valuable information about their evolutionary
history. The genomes of these two species are nearly identical, with
differences in only about 1-2% of their DNA. This suggests that
humans and chimpanzees are closely related, and that they have
evolved from a common ancestor.
96
Interrelatedness also has practical applications in medicine and
biotechnology. By comparing genomes, scientists can uncover
genetic causes of diseases and develop new treatments.
Three types of genomes can be compared:
1.
Nuclear genome: This refers to the genetic material found in
the nucleus of a cell, which includes both the chromosomes
(structures that carry the genetic information) and the nonchromosomal DNA. In most organisms, the nuclear genome
is the largest and most complex component of the genome.
2.
Mitochondrial genome: This refers to the genetic material
found in the mitochondria, the organelles that produce
energy for the cell. The mitochondrial genome is much
smaller than the nuclear genome and typically contains only
a few genes that are important for energy production.
3.
Chloroplast genome: This refers to the genetic material
found in the chloroplasts, the organelles that carry out
photosynthesis in plants and algae. The chloroplast genome
is also smaller than the nuclear genome and contains genes
involved in photosynthesis and the maintenance of the
chloroplast.
The degree of relatedness between genomes can be determined by
comparing their genetic sequences. This can be done at several
levels, including:
1.
DNA sequence: This refers to the specific order of
nucleotides (A, C, G, and T) in a DNA molecule. Comparing
DNA sequences can reveal similarities and differences
between genomes and can provide information about
evolutionary relationships.
97
2.
Gene content: This refers to the number and types of genes
present in a genome. Comparing gene content can reveal
similarities and differences between genomes and can
provide information about the functions and adaptations of
different organisms.
3.
Chromosomal structure: This refers to the organization and
arrangement of chromosomes within a genome. Comparing
chromosomal
structure
can
reveal
similarities
and
differences between genomes and can provide information
about the evolutionary history of different organisms.
Overall, the degree of relatedness can vary greatly between
genomes, with some organisms being very closely related (such as
different species within the same genus) and others being more
distantly related (such as different phyla or kingdoms).
Principle
The process of measuring the similarity of DNA sequences from
different species is known as hybridization. This technique is used to
estimate the extent to which organisms from different species have
common DNA sequences.
The process of hybridization is carried out in a series of steps. First, a
solution of denatured DNA from one species is filtered through a
membrane. The single-stranded DNA sticks to the filter and becomes
irreversibly bound to it when the filter is air dried.
Next, the filter is incubated in a solution of denatured heterologous
DNA. The incubation is done at a temperature 25°C below the
melting temperature of the denatured DNA in the solution. The
incubation causes duplexes to form between the filter-bound DNA
and the DNA in the solution. If the DNA in the solution is isotopically
labeled, the hybrids will bind the label to the filter. However, free
98
hybrid molecules are formed in the solution but they are not
detected as they are not bound to the filter.
The experiment measures the differential ability of some heterologous DNA to form DNA / DNA hybrids, thus estimating the
similarities of the different DNA sequences. This is done by
comparing the amount of hybrids formed when no competitor is
present versus the amounts of hybrids formed in the presence of
different quantities of competitor. Before starting the experiment,
the DNA solutions are denatured by placing them in a boiling water
bath for 10 minutes.
Procedure
Step-1: Preparing the Tubes for DNA Analysis
1.
Collect tubes for each competitor's DNA.
2.
Add a homologous and heterologous DNA filter to each
tube.
3.
Gently tap the tubes to mix the contents.
4.
Add 0.5 ml of fluid to cover the top of the tubes.
5.
Cover the tubes and place them in a water bath set at 61°C
for 18 hours.
Step-2: Analyzing the Filters for Hybrid Formation
1.
An hour before the incubation period ends, place a rack
containing l XSSC in the water bath.
2.
Remove the filters from the tubes and transfer them to l
XSSC.
3.
Rapidly stir the filters in l XSSC.
4.
Transfer the filters from one row of tubes to the next, until
they are all in the third row.
5.
Blot the filters dry on a sheet of paper towel.
99
6.
Pin the filters to the towel and incubate at 60°C for 20
minutes.
7.
Place the filters in a scintillating vial and count for 10
minutes.
8.
Discard the filters into a container for radioactive waste.
Step-3: Data Treatment and Analysis
1.
Subtract the counts per minute of the heterologous filter
from the homologous filter.
2.
Express the results as percentages.
3.
Plot the results graphically to show the percent of hybrids
formed against the amount of competitor DNA added.
100
In situ Hybridization
In situ hybridization studies on chromosomes provide an approach
to genetic mapping of the sequence of interest. When one of the
hybridization partner remains in situ, using a given labeled
polynucleotide (DNA or RNA) probe, location of the homologous
sequences in cells can be determined. The pattern of functional
organization or its expression can also be studied conveniently by
this technique at cellular or at organ level.
In situ hybridization (ISH) is a laboratory technique used to detect
and map the location of specific RNA or DNA sequences within cells
or tissues. The process involves labeling a specific RNA or DNA
probe with a fluorescent or radioactive tag, and then hybridizing the
probe to its complementary target sequence within the sample.
The basic steps of ISH include:
1.
Sample preparation: The tissue or cells are fixed and
embedded in a paraffin block or frozen section.
2.
Probe preparation: The specific RNA or DNA probe is labeled
with a fluorescent or radioactive tag, such as digoxigenin or
biotin.
3.
Hybridization: The labeled probe is added to the sample and
allowed to hybridize to its target sequence. The probe will
only bind to its specific target sequence, allowing for the
detection and localization of that specific sequence within
the sample.
101
4.
Detection: The hybridized probe is detected using an
appropriate detection method, such as a fluorescent
microscope or a auto-radiography.
5.
ISH is a powerful technique that allows for the visualization
of specific RNA or DNA sequences within cells or tissues, and
can
be
used
to
study
gene
expression
patterns,
chromosomal abnormalities, and viral infections, among
other applications.
Stock
1.
2.
20X SSC
Sodium chloride
3M
Sodium citrate
0.3 M
Gelatin solution
0.1%
Gelatin
100 mg
Distilled water
100 ml
(warm at 70 C for 1 hr)
o
3.
Sodium acetate
3M
(pH 5.2 with the help of glacial acetic acid)
4.
RNase
10 mg /10 ml
5.
Alcohol 70/90/100 %
6.
Tris
(pH 7.5, pH 8.0, pH 9.5) 1 M
Sodium chloride
EDTA
5M
0.05 M
TE
7.
Tris
10 mM (pH 8.0)
102
EDTA
1 mM
8.
Digoxigenin-dUTP labeled DNA probe
9.
Salmon sperm DNA 10 mg/ml
10.
Hybridization mix
11.
Buffers
I
12.
-
Tris 100 mM, pH 7.5; NaCl 150 mM
II -
Blocking reagent in Buffer - I 0.5% w/v
III -
Tris 100 mM, pH 9.5 ; NaCl 100 mM, MgCl2 50 mM
IV -
Tris 10 mM, pH 8.0 ; EDTA 1mM
Developer (to be prepared fresh)
Nitroblue Tetrazolium 4.5 μl
(NBT 75 mg/ml in Dimethyl formamide)
5-Bromo-4-Chloro-3-Indolyl Phosphate 3.5 μl
(50 mg/ml in Dimethyl formamide)
Buffer - III to make 1 ml
13.
Stains
Safranin 100 mg
Distilled water 100 mg
(Dissolve Safranin powder in water at room temperature. The
prepared stain can be stored and used later on).
14.
Entellan mountant
Laboratory ware
1.
Three incubators, preset at 37o, 42o and 60oC.
2.
Sterilized glass slides and cover glasses
3.
Slide racks
4.
Slide trays
5.
Couplin jars
103
6.
Magnetic stirrer
7.
Micropipettes
8.
Pipette tips
9.
Plastic box
10.
Forceps
Procedure
Step 1: To label the probe DNA
Chromosome in situ hybridization is a method used for genetically
mapping a specific sequence of DNA. This method uses a labeled
polynucleotide (DNA or RNA) probe to identify the position of
homologous sequences in cells when one of the hybridization
partners is still in place. This technique allows for the study of the
pattern of functional organization or expression at the cellular or
organ level.
One way to label the probe DNA is through the Random Priming
Method, which involves the following steps:
1.
Denature a necessary quantity of linear DNA by heating it in
boiling water for 10 minutes, then quickly chilling it on ice.
2.
Add 2 μl of hexanucleotide mix, 2 μl of dUTP labeled mix, 19
μl of distilled water, and 1 μl of Klenow enzyme (3-5 units) to
the chilled DNA.
3.
Incubate the mixture at 37°C for an hour, then add 0.8 ml of
0.5 M EDTA to stop the process (final concentration 20m M)
4.
Add 2.0 μl of salmon sperm DNA (10 mg/ml), 2.5 μl of
lithium chloride (4 M), and 75 μl of prechilled (-20oC) ethanol
to the labeled DNA to precipitate it.
5.
Incubate the mixture at -70oC for two hours, then centrifuge
the tubes for 30 minutes at 4°C at 12,000 rpm.
104
6.
Separate the supernatant from the pellet and wash it in 70%
ethanol. Dry in a warm environment.
7.
Dissolve the labeled probe in the necessary volume of TE.
The labeled probe can be stored at -20oC for more than two years.
Step 2: To test the effectiveness of DIG labeling
1.
Take a small piece of nylon membrane and moisten it with
2X SSC while under vacuum. Allow it to dry at room
temperature for about 30 minutes.
2.
Cross-link the probe DNA with the membrane by either
heating the filter for two hours at 70°C or by exposing it to
UV for three to four minutes on a transluminator.
3.
Clean the filter briefly with Buffer I.
4.
Block the membrane surface for 30 minutes at room
temperature in Buffer II to facilitate the subsequent use of an
antibody that will bind specifically to it. Use Buffer-I to rinse.
5.
Incubate in Anti-DIG Antibody-enzyme conjugate (1 μl in 4
ml of buffer- I) for 30 mins at room temperature.
6.
Wash twice in Buffer I with a 15-minute gap between each
wash. Rinse briefly with Buffer- III
7.
Place the blot in a tiny polythene bag, add the color
developer, and then seal the bag in a dimly lit area.
8.
Wrap the blot with aluminum foil and incubate it inside a
sealed bag in a dark closet until the desired level of color
signal appears.
9.
Take the blot out of the bag and keep it in buffer IV to stop
the reaction once a sufficient signal arises. The blot should
be kept dry or in buffer IV.
105
Notes
Under ideal probe labeling circumstances, 0.1 pg of probe produces
a measurable signal in less than 30 minutes.
Step 3: To take care of prepared slides
1.
First submerge the slides in a freshly made 0.1% solution of
gelatin for 3-5 seconds. This is done to apply a coat of
gelatin on the slides which helps to keep the cells in a good
state by preventing the binding of the probe. After that, airdry the slides.
2.
Place 100 ml of RNase (100 g/ml in 2 X SSC) over the
material on the slides. Then, cover each preparation with 22
mm2 cover glasses and place the slides in a moist chamber
with filter papers soaked in 2 X SSC. The slides should be
incubated for 2 hours at room temperature. This step helps
to remove any RNA from the preparations.
3.
After the 2-hour incubation, gently dip the slides into a
beaker containing 2 X SSC and allow the cover glasses to fall
into the solution. Clean the slides three times in 2 X SSC for
5 minutes each, twice in 70% ethanol for 10 minutes each,
and once in 95% ethanol for 5 minutes. If necessary, air dry
and store the slides in that manner. This step helps to
remove any remaining RNase and clean the slides.
4.
Now, immerse the slides in 0.07 N-NaOH for 3 minutes to
denature chromosomal DNA. This step helps to break down
any DNA present on the slides.
5.
Finally, wash the preparations twice in 95% ethanol for 5
minutes each and three times in 70% ethanol for 10 minutes
each. Air dry the slides. This step helps to remove any
remaining NaOH and clean the slides. The prepared slides
are now ready for further use.
106
Hybridization mix
1.
Formamide
500 μl
2.
20 X SSC
250 μl
3.
DIG labeled probe (per slide)
10-20
4.
H2O to make 1000 μl
ng
*For one slide, 15 to 20 μl of hybridization mix is sufficient.
Step 4: Hybridization
Hybridization is a process used to identify specific regions of DNA on
a chromosome. The following steps outline the procedure for
performing hybridization using a tagged probe DNA and a
hybridization mixture.
1.
To denature the tagged probe DNA, place the tubes
containing the probe in a boiling water bath for 10 minutes.
This will cause the double-stranded DNA to separate into
single strands. After denaturation, add the desired quantity
of denatured probe to the hybridization mixture.
2.
Mix 20 μl of the hybridization mixture with 10-20 ng of the
labeled probe. Place a cover glass over the mixture and seal
the edges using DPX.
3.
Incubate the slides for 12-14 hours at 37°C in a humid,
closed environment. This will allow the probe to bind to its
complementary sequence on the chromosome.
4.
After the incubation period, remove the DPX sealing with
forceps and dip the slides in 2 X SSC to remove the cover
glass.
5.
The slides should be washed three times in one SSC for 15
minutes each at 60°C. This step is important to remove any
unbound probe and other contaminants.
107
Step 5: Detection of Color
1.
Rinse the slides for one minute in Buffer I and 30 minutes in
Buffer II. This step is necessary to remove any remaining
contaminants from the previous washing step.
2.
Rewash the slides for one minute in Buffer I.
3.
Incubate the slides in anti-Digoxigenin antibody alkaline
phosphatase conjugate for 30 minutes after diluting it
1:5000 in buffer I. This step is necessary to bind the probe to
the chromosome.
4.
Wash the slides twice for two minutes each in Buffer I,
followed by a rinse in Buffer III.
5.
Depending on the desired signal, place 20-30 μl of freshly
made color reagent on the slide, cover it with a cover glass,
and seal it with DPX. Leave the slide in a dark room at room
temperature for 1-12 hours.
6.
Examine the slides under a microscope before placing them
in Buffer IV to stop the reaction.
7.
After counterstaining with safranin for 5-10 seconds and airdrying, wash the surface twice with distilled water.
8.
Allow the finishing touches to air dry before mounting with
Entellan (E. Merk).
Result
The hybridized probe binds to specific regions of the chromosomes,
causing
a
purplish-blue
color
deposit.
By
using
polytene
chromosome maps, it's possible to identify the exact location of the
hybridization signal.
Notes
Several factors must be considered to ensure that the hybridization
signal is strong and accurate.
108
1.
The quality of the chromosomal preparation is decisive. If
the
chromosomes
are
not
prepared
properly,
the
hybridization signal may be weak or not visible at all.
2.
The process of denaturing the chromosomes must be
precisely regulated to avoid ruining the structural details.
The probe must also be denatured right before use.
3.
Inadequate washing after hybridization and
antibody
binding may result in unwanted background. It is important
to properly wash the slides to remove excess probe and
antibody.
4.
The use of safranin can improve the shape and contrast of
the chromosomes, making it easier to see the hybridization
signal. If safranin is not available, an alternative stain 2%
aceto-orcein can be used. However, care must be taken to
prevent the staining from masking the hybridization signal.
5.
It is important to avoid air bubbles while mounting the cover
glass. Air bubbles can impede the local response and
obstruct the hybridization signal, leading to inaccurate
results.
109
Amplification of DNA using
Polymerase Chain Reaction
Polymerase chain reaction, commonly known as PCR, is a laboratory
technique used to amplify specific DNA sequences. It involves
multiple steps, starting from identifying the target sequence,
designing and checking the specificity of the primers, optimizing the
PCR conditions, analyzing the results, and finally, starting the
reaction and visualizing the results.
Primer Design
Primer design is an important step in the PCR process. Effective
primers must be designed to ensure accurate amplification of the
target sequence. The primer sequence must be complementary to
the flanking sequences of the target region and should not contain
any repeat or run sequences that could cause mispriming. The
primer should match the target sequence at the 3’ end, and there
should not be any complementary sequences between the primers
to prevent primer dimers. The primer length should be between 1825 base pairs and the optimal GC content should be between 4060%. The GC content is calculated by dividing the number of G and C
bases by the total number of A, T, G, and C bases and multiplying by
100.
Melting temperature (Tm) is a measure of the temperature at which
the double-stranded DNA molecule will separate into two single
strands. It is important for the Tm to be within the range of 50-60°C
and for the Tm difference between the forward and reverse primer
110
not to exceed 5°C. The Tm can be calculated using the formula [(G +
C) * 4] + [(A + T) * 2].
Annealing temperature (Ta) is the temperature at which the primers
bind to the target DNA. It is important that the Ta is not too high or
too low, as this can lead to low product yield or non-specific
products. The presence of G or C bases within the last five bases
from the 3' end of primers is known as GC clamp, and it promotes
specific binding at the 3' end. The GC clamp should not be more
than 2 Gs or Cs.
GC clamp refers to the presence of G or C bases within the last five
bases from the 3' end of primers. It promotes specific binding at the
3' end and should not be more than 2 Gs or Cs. There are various
tools available for primer design, such as the NCBI Primer design
tool and Primer3 or primer3Plus, and it is important to check for
specificity by running a BLAST on NCBI.
PCR optimization is the process of finding the most efficient set of
conditions for a PCR reaction. Different reactions may require
different conditions to yield the best results, and it is important to
optimize the conditions for the best results.
Post-PCR analysis refers to the analysis of the products of a PCR
reaction, known as amplicons. One common method of analysis is to
separate the DNA by size using an agarose gel. The gel provides
visual evidence of the success or failure of the reaction and the
concentration of agarose used is determined by the size of the
amplicon.
PCR has several advantages, including simplicity, ease of use,
sensitivity, and availability of reagents and equipment. The standard
operating procedure for the technique has been extensively
validated.
111
PCR has many different applications, including genotyping, cloning,
mutation detection, sequencing, microarrays, RT-PCR, forensics, and
paternity testing. Its versatility makes it a valuable tool in many
different fields of research and analysis.
1.
To amplify a specific region of DNA.
2.
To prepare right primers.
3.
To determine the parameters that may affect the specificity,
fidelity, and efficiency of PCR.
Stock
1.
Purified genomic DNA of male and female Calotes
2.
Stock solution of each primer 10 pM/μl
3.
10X Taq Polymerase Buffer
KCl 500 mM
Tris-HCl (pH 8.4 at 26oC) 100
mM
MgCl2 15 mM
4.
Gelatin 1 mg/ml
5.
DNTPs mix
(each of the 4 dNTPs) 1.25 mM
6.
Mineral Oil
7.
1% Agarose gel
Laboratory ware
Thermal cycler/ 3 water baths set at 92oC, 72oC and 60oC.
Procedure
The process of amplifying DNA through Polymerase Chain Reaction
(PCR) requires careful selection of DNA using two primers. These
primers are designed to match two regions of potential sequence
conservation that are separated by a DNA segment of around 60 to
112
1000 nucleotides. The design of the PCR primers is a crucial aspect of
the amplification process.
To create the primers, the first step is to identify conserved protein
regions. Then, degenerate oligonucleotide primers are synthesized
to match the potential DNA sequences that could encode the amino
acids. To match the DNA sequences that could encode the Nterminal protein patch, a mixture of forward primers is synthesized.
These primers are positioned more towards the 5’ end of the gene.
On the other hand, reverse primers are positioned more towards the
3’ end of the gene, and are made to match the inverse complement
of DNA sequences that could encode the C-terminal protein patch.
When designing the primers, the following guidelines should be
followed:
1.
Each degenerate primer should roughly match the potential
encoding DNA sequences for at least 15 to 20 nucleotides.
2.
Degeneracy should be introduced into the 3’ half of the
primer sequence (at 6-10 nucleotide positions) in such a way
that all possible codons that could code for the conserved
amino acids are represented.
3.
The 3’ terminal base of the forward primer should match
either the second codon position for an amino acid encoded
by 2, 3 or 4 codons (excluding Arg, Leu, Ser) or the third
position of a Met or Trp codon. The 3’ terminal base of the
reverse primer should compliment the first base of a codon
specifying an amino acid encoded by few alternative codons
(e.g, Met, Cys, Trp, His....).
4.
Degeneracy should be kept low (up to 1024 fold) by proper
selection of priming sites.
113
5.
Primer sequences should not have mono or dinucleotide
polymers and should not end on three G or C residues at the
3’ end.
6.
If the genes you're trying to clone are more varied than the
species you have data for, design primers around regions
that are rich in Cys, His, and Pro. These amino acids are less
likely to change. Avoid regions rich in Ser, Ala, Asp, Glu, Phe,
Tyr, and Arg because they are more likely to change........
7.
If the amplified DNA fragment becomes longer than 600
base pairs, the 5’ end of each primer should be altered in
such a way that a six-cutter restriction site is present.
8.
Using a compatible software, ensure that the designed
primer sequences do not form stable hybrids with each
other
or
amongst
degenerate
primer
mixes.
Avoid
complementarity of more than three contiguous bases if
these lie at the extreme 3’ end of a primer.
Proper primer design is crucial for successful PCR amplification, and
the guidelines provided above can be used as a reference to ensure
that the primers will match the desired DNA sequences and produce
accurate results.
Procedure
1.
Take two clean Eppendorf tubes of 1.5 ml each (one for male
and another for female genomic DNA) and add …….
10x Polymerase Buffer 5 μl
dNTPs mix 8 μl
Primers (10 pM/ul) 3 μl each
Genomic DNA (male and female separately) 100 ng
each
Taq polymerase 1 U
Distilled water to make 50 μl
114
Mix and overlay with 50 ul of mineral oil.
2.
Set up the PCR machine with appropriate cycling conditions
(temperature, time, and number of cycles). For example, an
initial denaturation step at 95°C for 3 minutes, followed by
35 cycles of denaturation at 95°C for 30 seconds, annealing
at 55°C for 30 seconds, and extension at 72°C for 1 minute.
The final extension is usually carried out at 72°C for 10
minutes.
3.
To visualize the amplified DNA bands, load the PCR products
onto an agarose gel and run gel electrophoresis. Capture an
image of the gel using a gel imaging system.
Result
The polymerase chain reaction (PCR) amplifies the target DNA
sequence through a series of repeated cycles that involve
denaturation, annealing, and extension. The resulting PCR product is
a mixture of DNA fragments of varying sizes, which can be analyzed
using gel electrophoresis. The detection of a specific DNA band on
the gel confirms the successful amplification of the target sequence.
Advantages and Disadvantages
PCR is a powerful tool for isolating and amplifying specific DNA
sequences, making it ideal for disease diagnosis through genetic
mutation detection.
However, there are two main drawbacks associated with PCR:
1.
The use of DNA polymerase can sometimes lead to errors
and mutations in the amplified DNA.
2.
The primers used in PCR can sometimes bind incorrectly to
the template DNA, leading to the production of non-specific
PCR fragments.
115
Real time PCR
Procedure
Prepare the qPCR reaction mix in a sterile microcentrifuge tube:
•
Add 10 μl of qPCR buffer.
•
Add 0.5 μl each of primers (10 μM stock solution).
•
Add 0.5 μl of fluorescent probe (10 μM stock solution).
•
Add 0.25 μl of DNA polymerase with a fluorescent probe (5
U/μl stock solution).
•
Add 4.25 μl of sterile water.
•
Add 10 μl of DNA sample (concentration varies depending
on the source and quality of DNA).
Thoroughly mix the contents of the tube using a pipette.
Transfer the reaction mix to a qPCR tube or plate that is compatible
with the qPCR machine.
Set up the qPCR machine with appropriate cycling conditions
(temperature, time, and number of cycles), such as:
An initial denaturation step at 95°C for 3 minutes.
Followed by 40 cycles of denaturation at 95°C for 15
seconds, annealing/extension at 60°C for 60 seconds, and
plate read at 60°C.
Analyze the qPCR data using suitable software to obtain a
quantification cycle (Cq) value, which indicates the point at
which the fluorescence signal of the amplified DNA reaches
a threshold level. The Cq value is used to calculate the
amount of DNA present in the sample.
116
Result
Real-time PCR, also known as quantitative PCR (qPCR), enables realtime detection and quantification of amplified DNA during the
amplification process. This technique utilizes fluorescent dyes or
probes that specifically bind to the amplified DNA product, allowing
for real-time monitoring of the reaction. The results are usually
presented as a graph of fluorescence intensity over time, which
enables determination of the amount of starting DNA template and
the efficiency of the amplification reaction.
Note
Prior to conducting the experiment, it is essential to take
necessary precautions to ensure safety in the laboratory,
maintain the quality and purity of the starting DNA material,
optimize primer design and reaction conditions, and
minimize potential sources of contamination.
Accurate
and
reliable
results
can
be
ensured
by
incorporating appropriate positive and negative controls.
In real-time PCR, calculating the threshold cycle (Ct) value
with precision is critical. The Ct value indicates the point at
which the fluorescence signal reaches a set threshold level.
Using suitable software for data analysis is also crucial.
117
Optimization of Annealing
Temperature
•
To refine the parameters that impact PCR results.
•
To fine-tune the annealing temperature for optimal PCR
results.
•
To gain proficiency in the PCR process and the use of a
thermal cycler.
Principle
To achieve optimal PCR results, familiarity with both the PCR
technique and thermal cycler device is essential. This involves a
comprehensive understanding of the basic concepts of PCR,
including the functions of each reagent, the PCR reaction
mechanism, and the thermal cycling steps. Familiarity with the
thermal cycler device's specific features such as temperature range,
ramp rate, and programmable settings is also necessary to optimize
the PCR reaction.
Procedure
PCR Optimization
In order to optimize a PCR reaction, it's important to adjust and find
the best concentration of each of the components involved in the
reaction. Each component has a typical or optimal concentration
range, which needs to be determined through a series of
experiments. The goal is to find the optimal concentration for each
component to achieve the most efficient and specific amplification of
the target DNA. It's important to note that while optimizing one
118
component, the concentrations of other components should remain
constant to avoid interference. These components include the Taq
polymerase, deoxyribonucleotides (dNTPs), magnesium, forward and
reverse primers, and the DNA template. To determine the ideal
concentration, optimization should be done one component at a
time, while keeping the other components constant.
The optimal concentrations of these components are generally
within the following ranges:
Taq polymerase at 1.25 units
dNTPs at 200 μM each
magnesium at 1.5-2.0 mM
primers at 0.1-0.5 μM each
DNA template at 1ng-1μg
The aim of optimizing the component concentrations is to find the
combination that provides the best reaction conditions for a
successful PCR.
Thermal Cycle
Thermal cycling optimization is the process of finding the best
temperature and duration for each step in the Polymerase Chain
Reaction (PCR) process. The objective is to obtain the optimal results
by determining the best temperature, duration and number of cycles
for each step.
There are three stages in the thermal cycling process. The first stage
is the initial denaturation which takes place at a temperature range
of 94-97°C and lasts for 3 minutes. The purpose of this step is to
denature the template and activate the DNA polymerase.
The second stage is where the PCR process is repeated 25 to 35
cycles. This stage consists of three steps, denaturation, annealing,
119
and elongation. Denaturation occurs at 94-97oC and lasts for 30
seconds. Annealing takes place at a temperature range of 50-65oC
and lasts for 30 seconds. The elongation step occurs at 72-80 oC and
lasts for 30 seconds to 1 minute.
The final stage is the final elongation phase which lasts for 5-7
minutes. This step is crucial as it allows for the synthesis of many
uncompleted amplicons to finish.
Annealing Temperature
The optimal annealing temperature refers to the temperature that
allows the primers to bind best to the target DNA. Finding the
optimal temperature is crucial to ensure that only specific products
are produced and non-specific products are avoided. A common
approach to optimize the annealing temperature is to incrementally
raise or lower the temperature in small steps and measure the
amount of amplified product produced at each step, until the
optimal temperature is found.
The temperature gradient PCR technique is often used to find the
optimal annealing temperature. This technique involves performing
the PCR reaction at different temperatures starting from 5°C below
the calculated melting temperature of the primer pair. For instance, if
the melting temperature of the primer pair is 58°C, the annealing
temperature will start from 53°C and will be increased by 8 degrees,
typically ranging from 53-60°C.
Stock
1.
PCR buffer
2.
DNA Taq polymerase
3.
dNTPs
4.
MgCl2
5.
Primers
120
6.
DNA template
7.
Nuclease free water
Procedure
To carry out a PCR reaction
Step 1: Determine the standard concentration of PCR components:
1.
Prepare a table to calculate the volume of each component
needed.
2.
Components include PCR buffer, Taq polymerase, dNTPs,
MgCl2, forward primer, reverse primer, DNA template, and
water
Step 2: Prepare the master mix
1.
2.
Mix all the components except for the DNA template.
Multiply the volume of each component by the number of
desired reactions plus one to account for pipetting error.
3.
Distribute the master mix into special PCR tubes using
pipettes.
4.
Add the DNA template to each tube.
Step 3: Centrifuge the tubes
Briefly spin the tubes to ensure that the components are well
mixed
Step 4: Set the thermal cycling conditions
1.
Temperature, time, and number of cycles must be set
2.
Different annealing temperatures may be tried based on the
primer pair Tm
Step 5: Start the PCR reaction
The final volume in the thermal cycler must be set to 50 μl
121
Result
1.
Use a 2% agarose gel to analyze the results
2.
Determine the optimum Ta.
122
Reverse Transcription PCR | RT-PCR
Reverse transcription polymerase chain reaction (RT-PCR) is a
technique that combines the principles of reverse transcription (RT)
and PCR to amplify and detect specific RNA sequences. Reverse
transcription is a process that converts RNA into complementary
DNA (cDNA) using an enzyme called reverse transcriptase. PCR is a
technique that amplifies specific DNA sequences using the enzyme
polymerase and a set of specific primers.
The basic steps of RT-PCR are as follows:
1.
Reverse transcription: The RNA sample is mixed with reverse
transcriptase, a primer (often called an oligo-dT primer) that
binds to the poly-A tail of mRNA, and other reagents such as
dNTPs (deoxynucleoside triphosphates) and MgCl2. This
mixture is then heated to allow the reverse transcriptase to
synthesize cDNA from the RNA template.
2.
PCR amplification: The cDNA is then used as a template for
PCR amplification. The PCR reaction mixture contains the
cDNA, a set of primers that specifically bind to the target
sequence of interest, and the polymerase enzyme. The PCR
reaction is typically performed in a thermal cycler, where the
sample is repeatedly heated and cooled to allow the primers
to bind to the template and the polymerase to synthesize
new DNA strands.
123
3.
Detection: The amplified PCR product can then be detected
by
various
methods,
such
as
gel
electrophoresis,
fluorescence, or sequencing.
RT-PCR is a highly effective method that enables the detection and
quantification of specific RNA sequences, even in very small
amounts. This technique is used across various fields such as
molecular biology, genetics, and medicine. It serves various purposes
such as analyzing gene expression, identifying infectious diseases,
and discovering cancer markers.
The technique is frequently utilized in gene expression profiling, to
evaluate the level of gene expression and determine the sequence of
an RNA transcript. It can also be used to identify the position of
exons and introns when the genomic DNA sequence of a gene is
known. To identify the 5' end of a gene, which represents the
starting point of transcription, RACE-PCR (Rapid Amplification of
cDNA ends) is performed.
When studying gene expression during development, two main
aspects are considered, whether a specific gene is expressed in an
embryo and where is the gene expressed in the embryo. Techniques
such as RNA extraction and northern blotting can demonstrate
expression, but in situ hybridization is needed to determine the
specific location of expression. However, low levels of gene
expression can make these techniques difficult to perform, so RTPCR is used to amplify transcripts for analysis and cDNA cloning.
One of the main advantages of RT-PCR is its sensitivity, which allows
for the detection of low-abundance RNA molecules. Besides, RT-PCR
can be used to detect both known and unknown sequences, making
it a valuable tool for discovery-based research. However, RT-PCR also
has some limitations, such as the potential for false positive results,
the need for specific primers, and the potential for contamination.
124
Stock
1.
Stock solution - D
Guanidium isothiocyanate 10 gm
Water 11.72 ml
Sodium citrate (pH 7.4) 0.704 ml
10% Sacrocyl 1.05 ml
2.
Working solution - D
Stock solution - D 1 ml
2 – mercaptoethanol 7.5 ml
Sodium citrate (pH 7.0) 0.75 M
3.
10% N-lauryl sarcosine
4.
Water saturated phenol
5.
Chloroform
6.
Isopropanol
7.
Ethanol
8.
10 X DNase I reaction buffer
Sodium acetate 1 M
MgSO4 1 M
DNase I (RNase free)
9.
RNase inhibitor
EDTA 20 mM
Oligo dT 0.5 μg/ml
dNTP mix 10 mM
DTT 0.1 M
10.
10 X Synthesis buffer
11.
Reverse transcriptase
12.
Primers
13.
Taq DNA polymerase
125
14.
Agarose
(Solutions for RNA extraction should be DEPC treated)
*Glassware should be acid treated autoclaved and baked at 300oC
for 4 hrs and plastic wares should be rinsed with chloroform to
inactivate RNase.
Procedure
Step 1: Isolation of RNA by AGPC Technique
1.
Take out 10 mg of tissue in chilled PBS.
2.
Mince the tissue on an ice-slab and homogenize it with 100
μl of solution D. Place the contents in a test tube.
3.
Then, add 10 μl of 2M Sodium acetate (pH 4.0), 100 μl of
phenol, and 200 μl of chloroform to the test tube. Mix well in
a cyclomixer and store the mixture on ice for 15 minutes.
4.
After 15 minutes, centrifuge the sample at 10 K for 20
minutes.
5.
Carefully remove the aqueous phase (RNA), avoiding the
organic and interphase (proteins and DNA).
6.
Mix an equal volume of isopropanol with the RNA and store
it at -70oC for 4 hours.
7.
After 4 hours, centrifuge the RNA at 10 K for 20 minutes.
Discard the supernatant and dissolve the pellet in 25 μl of
solution D (1/4th volume).
8.
Transfer the suspension to an Eppendorf tube and
precipitate the RNA with 1 volume of isopropanol or 2
volumes of ethanol at -20oC for 4 hours.
9.
Centrifuge the RNA at 15 K for 10 minutes, while
maintaining the temperature at 4oC.
10.
Wash the pellet in 80% ethanol, sediment it and let it dry.
11.
Treat the extract in DEPC water at 65oC for 10 minutes, if
needed. If not, dissolve it in 0.5% SDS.
126
12.
Finally, store the RNA at -70oC for future use.
Step 2: DNase I Treatment to RNA samples
Prepare a work area with an ice slab. Gather the following materials:
1 μg of RNA, 1 RNase free microcentrifuge tube (0.5 ml), 1 μl of 10 X
DNase I reaction buffer, 1 μl of RNase inhibitor, 1 unit of
amplification grade DNase I, DEPC treated water, 1 μl of 20 mM
EDTA.
1.
Take 1 μg of RNA and transfer it to the RNase free
microcentrifuge tube.
2.
Add 1 μl of 10 X DNase I reaction buffer and 1 μl of RNase
inhibitor to the tube with RNA.
3.
Add 1 unit of amplification grade DNase I to the reaction
4.
Bring the volume to 10 μl with DEPC treated water.
5.
Place the tube with the reaction mixture on the ice slab and
mixture.
incubate it at 37°C for 15 minutes.
6.
After 15 minutes, inactivate DNase I by adding 1 μl of 20
mM EDTA to the reaction mixture and heating it at 65°C for
10 minutes.
7.
After 10 minutes, extract the mixture with phenol:chloroform
and then with chloroform only.
8.
Precipitate the mixture by adding 2 volumes of alcohol.
9.
Wash the mixture with 80% alcohol.
10.
Dry the mixture and dissolve it in DEPC treated water.
Step 3: Preparation of First cDNA Strand
1.
Take an autoclaved microcentrifuge tube and add 1-5 ug of
total RNA in 13 μl of DEPC treated water.
2.
Now, add 1 μl of oligo dT (0.5 mg/ml) to the tube and mix
gently.
127
3.
Heat the mixture to 70oC for 10 minutes and incubate on ice
for a minute.
4.
To the mixture, add the following components in order:
10 X synthesis buffer (2 μl)
dNTP mix (10 mM, 1 μl)
DTT (0.1 M, 2 μl)
RTase (200 U/μl, 1 μl)
5.
Mix all the components gently and collect the reaction
mixture after a brief spinning.
6.
Incubate the mixture at room temperature for 10 minutes.
7.
Transfer the tube to a water bath preset at 42oC and let it
stand for 50 minutes.
8.
Finally, terminate the reaction by increasing the temperature
to 70oC for 15 minutes and then cooling on ice.
Step 4: Polymerase chain reaction
1.
Prepare the reaction mixture.
Take a small sample of 1st strand cDNA (approximately 1 μl).
Add 8 μl of 10X synthesis buffer to the sample.
Add 1 μl of Primer 1 (10 μM) to the mixture.
Add 1 μl of Primer 2 (10 μM) to the mixture.
Add 1 μl of Taq DNA polymerase (5 U/μl) to the mixture.
Add water to the mixture to make a total volume of 80 μl.
2.
Overlay with mineral oil: Gently stir the reaction mixture and
add 2 drops (approximately 100 μl) of mineral oil on top to
prevent evaporation during heating.
3.
Initial denaturation: Place the reaction mixture in a
thermocycler and heat it at 94oC for 5 minutes to denature
the DNA.
4.
Repeat the PCR cycle 15-30 times:
128
Denature the DNA at 94oC for 1 minute.
Anneal the primers to the DNA at 50oC for ½ minute.
Synthesize the new DNA strand using Taq polymerase at
72oC for 1 minute.
5.
Gel electrophoresis: Remove 10-12 μl of the amplified DNA
from the reaction mixture and analyze it on an agarose gel
to confirm successful amplification.
Notes
The concentration of Mg++ may affect the PCR results and may need
to be optimized for different PCR conditions.
129
Digestion of DNA with RE
Restriction endonucleases are a type of enzyme found in bacteria
and other prokaryotes that have the ability to recognize and cut
specific sequences of nucleotides in double-stranded DNA. These
enzymes play a role in defending the cell against invading viral
bacteriophages by cleaving their DNA and preventing replication.
There are over 300 known restriction enzymes, and each is named
based on the organism it was isolated from.
When restriction enzymes cut DNA, they produce fragments called
restriction fragments that can have either a blunt end or a sticky end.
Both types of cuts are useful in molecular genetics and can be used
to join DNA fragments. Restriction enzymes play a significant role in
several areas of genetic research. These enzymes are designed to
help create new DNA molecules by cutting and splicing existing DNA
sequences through a process called recombinant DNA technology.
This allows scientists to produce genetic material with specific traits.
Restriction enzymes are also used to study the genetic structure of
DNA fragments and entire genomes. By cutting DNA into smaller
pieces, they can map out the genetic makeup of an organism and
provide valuable information for scientific research. This process is
known as Restriction Fragment Length Polymorphism (RFLP) and is
used to detect variations in the sequence of nucleotides in the
similar fragments.
130
Principle
The process of cleavage by restriction endonucleases (RE) involves
the incubation of genomic DNA or DNA fragments that have been
amplified using PCR. The RE enzyme is used to restrict the DNA at
specific sequences that it recognizes. This process results in the
production of fragments of different sizes, which can then be
separated using agarose gel electrophoresis. For instance, the RE
enzyme - MstII used here, cuts the DNA at the sequence '5CCTNAGG-3'. To ensure successful cleavage, the appropriate
conditions of temperature, pH, and ionic strength must be
maintained during the incubation process.
Stock
1.
DNA solution (0.5 μg/μl)
2.
MstII (3U/ μl)
3.
10X restriction buffer
4.
NaCl solution
5.
Nuclease free water
6.
0.5 M EDTA
Procedure
1.
Gather all necessary materials including a clean microcentrifuge tube, a DNA solution with a concentration of 1
μg/μl, 10X restriction buffer, a solution of NaCl, and water.
2.
Label the micro-centrifuge tube.
3.
Add the following components to the tube:
1 μl of the DNA solution
2 μl of the 10X restriction buffer
1 μl of the NaCl solution
15 μl of water
131
4.
Add MstII to the reaction mixture. The amount of MstII
should be 3 units for each microgram of DNA.
5.
Incubate the reaction mixture for 20 minutes at 37°C in an
incubator.
6.
Stop the reaction by adding 0.5 μl of 0.5 M EDTA.
7.
Add 5 μl of gel loading buffer to the reaction mixture.
This will allow the mixture to be loaded onto the gel and run through
electrophoresis to visualize the restriction enzyme's effects on the
DNA.
132
Digestion of DNA with RE in
Bacteriophage
Restriction enzymes are specialized proteins that are found in various
bacteria and single-celled organisms. These enzymes are designed to
search a length of DNA for a specific base sequence. This sequence
is usually between 4 and 6 base pairs long and is referred to as the
recognition site. When the enzyme finds this recognition site, it binds
to the DNA molecule and cleaves each of the double helix strands,
resulting in the fragmentation of the DNA molecule.
In this experiment, we will be working with a virus called
Bacteriophage λ. This virus is specially designed to infect bacteria
and has been the subject of many studies in molecular biology. We
will take the DNA from this small virus and cut it into smaller pieces
using restriction enzymes. The genome of Bacteriophage λ is quite
small, only containing 48,502 base pairs, which is much shorter than
the human genome, which contains about 3 billion base pairs.
After cutting the DNA into smaller pieces, we use electrophoresis to
separate the fragmented DNA on an agarose gel. This is
accomplished by adding a loading buffer to the DNA sample to
inhibit the restriction enzyme, and then exposing the sample to an
electric field overnight. This drives the DNA fragments to migrate
into the gel. Once the migration is complete, the gel is dyed with
methylene blue, which allows the DNA bands to become visible. The
gel can then be photographed and the pattern so received can then
133
be compared to a predicted result to figure out the location of
specific genes and regions within the genome.
1.
To learn about the nature and operation of a DNA restriction
enzyme.
2.
To gain proficiency in using micropipettes
3.
To become familiar with DNA electrophoresis
4.
To identify a DNA sample using a restriction digestion map
5.
To compare the λ DNA bands on a gel with a known λ DNA
restriction map
Stock
Equipment
1.
Electrophoresis chamber
2.
Container with TBE solution (1X)
3.
Cooler with crushed ice
4.
Freezer (frosty, if possible)
5.
Microtube rack
6.
Four microtubes
7.
Camera, if desired
8.
Gloves
9.
37°C water bath w/ floating rack
10.
60°C water bath or saucepan on a hot plate
11.
20-μl micropipette (or 10-μl micropipette) and sterile tips
12.
Waterproof pen
13.
500-ml beaker
Reagents
1.
Beaker or foam cup with crushed ice for the following:
20 μl of 0.4 μg/μl λ DNA
2.5 μl BamHI restriction enzyme
134
2.5 μl EcoRI restriction enzyme
2.5 μl HindIII restriction enzyme
10 l distilled water
2.
1.0% agarose gel
3.
20 μl 10X loading dye
4.
0.002% methylene blue stain
5.
Distilled water
Procedure
Step 1: Preparation for Gel Electrophoresis
Advance Preparations
1.
Check if the 1X TBE solution from the previous Gel
Electrophoresis with Dyes activity is still available for reuse.
2.
Collect enough ice cubes and foam cups for each lab group
to use as containers for keeping the restriction digests cool
during the lab.
3.
Heat a pan of water to 55°C on a hot plate to be used for
heating the restriction digests.
4.
Fill another pan with water and heat it to 37°C on a hot plate
to be used for heating the lambda DNA.
5.
To reconstitute the lambda DNA, add it to sterile distilled
water to reach a concentration of 0.4 g/l.
6.
For each group, measure out the necessary amounts of
lambda DNA, enzymes, and loading dye, and store them in
the freezer until the lab.
7.
Prepare the 1.0% agarose gel solution by melting 1.0 g of
agarose in 100 ml of 1X TBE buffer either in a microwave or
on a hot plate, and store it in the refrigerator if not in use
within the next 30 minutes.
135
To prepare gel for electrophoresis, follow these steps:
1.
Before starting the procedure, put on gloves and keep all the
necessary enzymes and DNA aliquots on ice to maintain
their integrity.
2.
Label 4 microtubes with the following labels - 10X buffer,
DNA, BamHI, EcoRI, HindIII and Water. Place these tubes in a
tube rack for ease of access.
3.
Using a micropipette set to 4 μl, add 4 μl of 10X buffer to
each of the four tubes. Make sure to use a new tip for each
buffer to avoid contamination.
4.
Using the same micropipette, add 4.0 μl of DNA to each of
the four tubes, again using a new tip for each sample.
5.
In the control tube, add 32.0 μl of distilled water and in the
other reaction tubes, add 30.0 μl.
6.
Close the microtubes and place them in a 55°C waterbath for
10 minutes to heat the samples. Immediately after, place the
tubes on ice for 2 minutes.
7.
Add 2 μl of the relevant restriction enzyme to each of the
reaction tubes. Make sure to use a new tip for each enzyme
to prevent contamination.
8.
Close the microtube caps and tap the bottom of the tubes
gently on the desktop to ensure all the liquid has settled at
the bottom. Finally, incubate the tubes overnight at 37°C.
The tubes will be frozen until use, and can be used within 60 days.
Step 2: Setting Gel on the Tray
1.
Heat a pan of water to 60°C.
2.
Pour enough agarose gels into the pan to warm and liquefy
them.
3.
Secure the ends of the gel tray with labeling tape and place
the plastic comb in the slots.
136
4.
Pour approximately 35-40 ml of agarose into each gel tray to
create a thick gel.
5.
Allow the gel to cool and solidify (about 15 minutes).
6.
Store the gel trays overnight in a container or ziploc baggie
with 0.5X TBE solution to prevent drying out.
Procedure for Setting Gel on the Tray and Loading buffer:
1.
Put on gloves. Fill a styrofoam cup with ice and place your
DNA digestion tubes on it.
2.
Take the 1.0% agarose gel and place it in the gel box with
the wells at the negative end.
3.
Carefully add 150 ml of 1X TBE solution to the gel box,
making sure the gel is covered with 2 mm of buffer.
4.
Gently remove the comb and make sure the buffer covers
the gel.
5.
Heat the microtubes in a 60°C water bath for 3 minutes to
ensure the DNA is in linear form.
6.
Use a micropipette set to 4 μl and add 4 μl of loading dye to
the bottom of each microtube.
7.
Set up the electrophoresis apparatus. Using a micropipette,
load 20 μl of each sample into a well.
8.
Turn on the power supply for 30-45 minutes and turn off
once the purple dye is 1 cm from the end of the gel.
9.
Unplug the gel box. To visualize the bands, place the gel in a
0.002% methylene blue solution in 0.1X TBE and stain
overnight at 4°C or for 2 hours at room temperature.
Step 3: Observation
1.
Remove the student gels from the refrigerator.
2.
Set up containers for staining near a sink and note that gels
can be discarded in the regular trash receptacle.
137
Proceed as follows:
1.
Take the gel and place it under white light.
2.
Look closely at the gel to see if the bands are visible.
3.
If the bands are not visible due to high background staining,
take a container filled with 0.1X TBE solution.
4.
Place the gel in the solution and agitate it gently.
5.
Change the buffer every 30 to 60 minutes. Continue this
process until the gel has reached the desired degree of destaining.
6.
If you wish, take a picture of the gel.
Restriction enzymes are special types of enzymes that can cut DNA
at specific locations. These locations, known as restriction sites, are
determined by the sequence of bases in the DNA, which often form
a repeating pattern that reads the same forwards and backwards.
These repeating sequences, called palindromes, are found on both
the forward and reverse strands of the DNA.
Restriction enzymes recognize and bind to these palindromic
sequences and then cut the DNA between specific bases. In the
present instance, there are three restriction enzymes used - EcoRI,
HindIII, and BamHI, each of which has its own unique recognition
sequence.
To measure the size of each of the fragments produced when λ DNA
is cut with each of these restriction enzymes, the fragments are
separated using electrophoresis and compared to a molecular
138
ladder. The molecular ladder has bands of known sizes and is
separated at the same time as the digested λ DNA. This allows for
the determination of the sizes of the fragments produced by each
restriction enzyme.
Notes
1.
Enzymes, particularly restriction enzymes, should be stored
care-fully and at a specific temperature to maintain their
activity.
2.
When working with Lambda (λ) DNA, it is recommended to
heat the sample to break the hydrogen bonds that hold it in
a circular form.
3.
Methylene blue dye is a less sensitive alternative to ethidium
bromide, but it can be used to stain larger quantities of
DNA. It should be handled with caution, as it can stain
clothes and equipment.
4.
When de-staining gels, only use distilled or deionized water
and make sure to wash the work area thoroughly.
5.
When making 0.1X TBE buffer, only use deionized water to
avoid damaging the DNA with high chlorine levels in tap
water.
139
Southern Blotting of DNA
Southern blotting is a technique used to detect specific DNA
sequences in a sample. The technique, named after Prof EM
Southern in 1975, involves the separation of DNA fragments by size
through electrophoresis, followed by transfer of the separated DNA
onto a solid support, such as nitrocellulose or nylon membrane. The
transferred DNA is then exposed to a labeled probe that binds to the
specific sequence of interest. The probe can be radioactive,
fluorescent, or enzyme-linked, and is usually complementary to the
target sequence. The probe-bound DNA can then be visualized using
autoradiography (for radioactive probes), fluorescence microscopy
(for fluorescent probes), or chemoluminescence (for enzyme-linked
probes).
The Southern blotting process can be divided into three main steps:
1.
Restriction enzyme digestion: The DNA sample is first
treated with one or more restriction enzymes to cut the DNA
at specific sites. This produces fragments of DNA of varying
sizes.
2.
Electrophoresis: The restricted DNA fragments are then
separated by size through electrophoresis. This is typically
done by running the fragments through an agarose gel,
which acts as a sieve to separate the fragments based on
their size.
3.
Transfer and hybridization: The separated DNA fragments
are then transferred from the gel to a solid support, such as
140
nitrocellulose or nylon membrane. This is done by a process
called blotting, which can be done by capillary action or by
electro-blotting. The transference of DNA from the gel to
the membrane involves:
Depurination: The agarose gel containing DNA is treated
with 0.2N HCl to depurinate the fragments and transfer
fragments larger than 8kb.
Denaturation: The denaturation solution denatures the
double-stranded DNA to single-stranded DNA, allowing
hybridization with the probe.
Neutralization: The neutralizing solution adjusts the pH to
enable hybridization.
Once the DNA is transferred, it is fixed in place on the membrane.
The membrane is then exposed to a labeled probe that binds to the
specific sequence of interest. The probe can be radioactive,
fluorescent, or enzyme-linked, and is usually complementary to the
target sequence. The probe-bound DNA can then be visualized using
autoradiography (for radioactive probes), fluorescence microscopy
(for fluorescent probes), or chemiluminescence (for enzyme-linked
probes).
Stock
Nylon membrane
Vacuum transfer apparatus
Shaker apparatus
UV Transilluminator
Micropipette
Agarose gel electrophoresis
141
Procedure
1.
Cut a nylon membrane slightly larger than the gel and
activate it in distilled water for 5-10 minutes.
2.
Set up the vacuum transfer apparatus and place the nylon
membrane, followed by the gel.
3.
Depurinate the DNA strand with 0.25N HCl.
4.
Wash the gel with distilled water and treat it with 0.5N
NaOH and 1.5N NaCl for 30 minutes.
5.
Neutralize the gel after denaturation by treating it with
neutralizing solution for 30 minutes.
6.
Maintain the vacuum pump and simultaneously pour buffer
over the gel using a pipette. Check the transfer carried out in
3 hours. After that, remove the gel and mark the pores on
the membrane with an HB-Pencil.
7.
Check the completion of transfer by viewing the nylon
membrane with a UV transluminator. Wash the membrane in
neutralizing solution, air dry, and bake at 80oC for 2 hours to
fix the DNA to the nylon membrane.
Result
The DNA smear on the membrane indicates successful transfer of
DNA. When the gel after transfer is viewed under UV light, no bands
are seen, indicating that the transfer is complete.
Advantages and Disadvantages
Southern blotting is a time-consuming and labor-intensive process
but it allows for the determination of molecular weight of restriction
fragments and the measurement of relative amounts of different
fragments in different samples.
142
Northern Blotting of RNA
Northern blotting is a technique used to detect specific RNA
sequences in a sample. The process is similar to Southern blotting,
which is used to detect specific DNA sequences. The main difference
is that Northern blotting is used to analyze RNA instead of DNA.
The Northern blotting process can be divided into three main steps:
1.
The RNA sample is first extracted from the tissue or cells of
interest. The RNA is then denatured by heating it in the
presence of formaldehyde or a similar chemical, which
causes the RNA to become single-stranded.
2.
The denatured RNA is then separated by size through
electro-phoresis. This is typically done by running the RNA
through an agarose gel, which acts as a sieve to separate the
RNA fragments based on their size.
3.
The separated RNA fragments are then transferred from the
gel to a solid support, such as nitrocellulose or nylon
membrane. This is done by a process called blotting, which
can be done by capillary action or by electroblotting. Once
the RNA is transferred, it is fixed in place on the membrane.
The membrane is then exposed to a labeled probe that
binds to the specific sequence of interest. The probe can be
radioactive, fluorescent, or enzyme-linked, and is usually
complementary to the target sequence. The probe-bound
RNA can then be visualized using autoradiography (for
radioactive probes), fluorescence microscopy (for fluorescent
probes), or chemo-luminescence (for enzyme-linked probes).
143
The Northern blotting technique is widely used in molecular biology
for detecting specific RNA sequences, for example for gene
expression analysis, studying alternative splicing, and for identifying
non-coding RNA. However, with the advancement of more sensitive
techniques like quantitative PCR (qPCR) and microarray, the use of
Northern blotting has declined. These newer techniques are more
efficient in detecting and quantifying RNA molecules, but Northern
blotting still holds its importance as it provides information about
the size of RNA, which could be useful in identifying degradation
products and determining the presence of specific isoforms.
Principle
The Northern blotting process involves using electrophoresis to
separate RNA samples based on their size and then using a
hybridization probe to detect the target sequence.
Procedure
1.
Obtain either total RNA or mRNA and prepare it for gel
electrophoresis.
2.
Load the RNA sample onto an agarose gel and run the gel
electrophoresis to separate the RNA molecules based on
size.
3.
Carefully transfer the separated RNA from the gel to a sheet
of nitrocellulose or other suitable blotting paper, ensuring
that the pattern of separation remains intact.
4.
Incubate the blot with a single-stranded DNA probe. This
probe will form base pairs with its complementary RNA
sequence and bind to form a double-stranded RNA-DNA
molecule. The probe can be radioactive or have an enzyme
bound to it (e.g. alkaline phosphatase or horseradish
peroxidase).
144
5.
Incubate the blot with a colorless substrate that the attached
enzyme can convert to a colored product that can be seen
or gives off light. Alternatively, if the probe was labeled with
radioactivity, expose X-ray film directly.
6.
Analyze the results to determine the size and quantity of the
RNA molecules present in the sample and the location of the
RNA-DNA hybrid.
Advantages and Disadvantages
Northern blotting has many benefits, such as being able to
determine the size of RNA, detect alternate splice products, use
probes with partial homology, and measure the quality and quantity
of RNA before blotting. It can also be used to study gene expression
by detecting specific RNA sequences in a mixture of RNA molecules.
Moreover, the membranes used in the process can be stored and
reused for years.
However, there are also some downsides to Northern blotting. Small
changes in gene expression may go undetected, and the samples
can be degraded by RNases. Compared to RT-PCR, Northern blotting
is less sensitive, but it has a higher specificity which reduces the
chances of false-positive results.
145
Western Blotting
Western blotting is a laboratory technique used to detect and
analyze specific proteins in a sample. The basic steps of the
procedure include:
1.
The sample, typically cells or tissue, is lysed to release the
proteins. The lysate is then separated by size using gel
electrophoresis. This creates a separation of the proteins
based on their molecular weight, with the smaller proteins
migrating further than the larger proteins.
2.
The separated proteins are then transferred from the gel to a
solid support, such as a nitrocellulose or PVDF membrane.
This step is called electroblotting.
3.
The membrane is then blocked to prevent non-specific
binding of the primary antibody. Common blocking agents
include bovine serum albumin (BSA) or non-fat dried milk.
4.
The primary antibody, which specifically binds to the protein
of interest, is then added to the membrane and incubated.
5.
A secondary antibody, conjugated to a detection enzyme or
fluorophore, is then added to the membrane. This antibody
binds to the primary antibody, thereby indirectly detecting
the protein of interest.
6.
The signal from the detection enzyme or fluorophore is then
visualized, typically using X-ray film or a digital imaging
system. The intensity of the band on the membrane
corresponds to the amount of protein present in the original
sample.
146
Western blotting is a common method used in the fields of
molecular biology, biochemistry, and medicine to detect and
quantify specific proteins in a sample of tissue homogenate or
extract. This technique helps researchers understand and describe
proteins, and can also be used to see how the expression levels of
proteins change when subjected to different stimuli or treatments.
Principle
The western blot technique involves the separation of proteins
through gel electrophoresis. The proteins can either be separated
based on their natural 3-D structure or their length if they are
denatured. After the separation, the proteins are transferred to a
membrane like nitrocellulose or PVDF and then stained with specific
antibodies to identify the target protein. The use of gel
electrophoresis is important in western blot analysis to eliminate any
potential issues with cross-reactivity of the antibodies.
Stock
Protein / antibody sample
SDS-PAGE gel
Nitrocellulose or PVDF membrane
Transfer buffer (such as Towbin's transfer buffer)
Blocking solution (such as 5% non-fat dry milk in TBS-T)
Primary antibody
Secondary antibody conjugated to a detection enzyme (e.g.,
HRP-conjugated
anti-rabbit
IgG),
Chemiluminescent
substrate (e.g., ECL substrate)
X-ray film.
147
Procedure
1.
Prepare an SDS-PAGE gel based on the desired molecular
weight range.
2.
Load the protein/antibody sample onto the gel and run the
gel according to the manufacturer's instructions.
3.
Transfer the separated proteins to a nitrocellulose or PVDF
membrane using a transfer buffer and transfer apparatus.
4.
Block the membrane with a blocking solution for 1 hour at
room temperature.
5.
Incubate the membrane with the primary antibody diluted in
blocking solution for 1 hour at room temperature or
overnight at 4°C.
6.
Wash the membrane with TBS-T buffer to remove any
unbound primary antibody.
7.
Incubate the membrane with the secondary antibody
conjugated to an enzyme for 1 hour at room temperature.
8.
Wash the membrane with TBS-T buffer to remove any
unbound secondary antibody.
9.
Incubate the membrane with a chemiluminescent substrate
for the required time.
10.
Expose the membrane to X-ray film and develop the film
according to the manufacturer's instructions.
Results
The Western blotting technique results in the appearance of a band
on the X-ray film, indicating the presence and amount of the target
protein in the sample. The technique is widely used in research and
diagnostic laboratories.
148
Notes
Handle the protein/antibody sample carefully to avoid
degradation or contamination.
Use appropriate safety precautions when handling chemicals
and X-ray film.
Ensure that the primary and secondary antibodies are
specific to the target protein and do not cross-react with
other proteins in the sample.
Other types of paper or membranes can be used in place of
nitrocellulose.
Advantages and Disadvantages
The western blot is a diagnostic tool that has several advantages and
disadvantages. The benefits of using western blot include its high
level of sensitivity and specificity in detecting anti-HIV antibodies, its
role as the definitive test for mad cow disease, and its use in some
forms of Lyme disease and Hepatitis B testing. In veterinary
medicine, it is also used to confirm the FIV status in cats.
However, the Western Blot is a complex and time-consuming
procedure
that
requires
specialized
equipment
and
trained
personnel. It can be expensive, especially when multiple tests are
required. In addition, the quality of the samples can impact the
results, leading to false negatives.
149
Dot Blot
Dot blot is a biochemical assay that is used to detect and quantify
the presence of specific target molecules in a sample. It is a simple,
quick and cost-effective method that can be used to detect a wide
range of targets, including DNA, RNA, proteins and small molecules.
In a dot blot, the sample is blotted onto a nitrocellulose or nylon
membrane and then probed with a labeled reagent, such as a
radioactively labeled antibody or DNA probe. The target molecules
on the membrane are then detected by autoradiography or
chemiluminescence, respectively. Dot blots are often used as a
preliminary screen for the presence of specific target molecules, or to
quickly test multiple samples for the presence of a specific target.
Principle
The principle of dot blot in biotechnology is to transfer a small
volume of sample onto a solid support such as nitrocellulose or
PVDF membrane, and then to detect specific biomolecules such as
DNA, RNA, proteins, or antibodies, using various detection methods
such as hybridization with labeled probes, immunodetection with
specific antibodies, or colorimetric reactions. The Dot Blot method
allows for quick and simple analysis of a large number of samples in
a small area, making it a useful tool for identifying specific molecules
in large numbers of samples. The method is commonly used in
applications such as DNA fingerprinting, protein identification, and
disease diagnosis.
150
Procedure
Dot blot is a method used to transfer nucleic acids or proteins from a
gel or liquid onto a nitrocellulose or nylon membrane. Here is a stepby-step guide to perform dot blot:
1.
The sample should be in a liquid form and can be extracted
from cells, tissues or bacteria.
2.
Cut the nitrocellulose or nylon membrane to the desired size
and place it on a flat surface.
3.
Using a micropipette, load the samples onto the membrane
in a small spot or dot form. Ensure that the dots are spaced
evenly on the membrane.
4.
Place the membrane on top of a vacuum manifold and apply
suction. This will cause the samples to be transferred from
the dots onto the membrane.
5.
Cross-link the samples to the membrane using a UV crosslinker or a chemical cross-linking agent. This helps to
prevent the samples from washing away during the
subsequent steps.
6.
To prevent non-specific binding, block the membrane with a
blocking buffer, such as 5% BSA. Incubate the membrane in
the blocking buffer for about 30 minutes.
7.
Incubate the membrane in the primary antibody for 1-2
hours. The primary antibody should be specific to the
protein or nucleic acid of interest.
8.
Wash the membrane with a washing buffer to remove any
unbound primary antibody. This step should be repeated
several times to ensure that all unbound antibody is
removed.
9.
Incubate the membrane in a secondary antibody that is
conjugated to a detection enzyme, such as horseradish
peroxidase (HRP).
151
10.
Wash the membrane again to remove any unbound
secondary antibody.
11.
Add a detection reagent, such as ECL or chemiluminescent
substrate, to the membrane. Incubate the membrane in the
detection reagent for 1-2 minutes. Visualize the result by
exposing the membrane
to X-ray film
or
using
a
chemiluminescent imaging system.
12.
The resulting dots will show the presence or absence of the
protein or nucleic acid of interest. The intensity of the dots
can be used to quantify the amount of the target molecule.
Advantages
1.
Dot blot is a simple and straightforward technique that does
2.
This technique is relatively cheap compared to other
not require specialized training or expensive equipment.
molecular
biology
techniques
such
as
PCR,
gel
electrophoresis, and Southern blotting.
3.
Dot blot is highly sensitive and can detect even trace
amounts of target molecules, making it ideal for detecting
low abundance analytes.
4.
Dot blot can be used to detect a variety of different types of
molecules such as proteins, DNA, and RNA.
5.
The dot blot technique is highly robust and can be
performed under a wide range of conditions.
Disadvantages
1.
One of the main limitations of dot blot is that it provides low
resolution compared to other techniques such as gel
electrophoresis.
2.
Dot blot can provide qualitative data but is not suitable for
accurate quantification.
152
3.
There is a possibility of interference from non-specific
binding or cross-reactivity, which can result in false positive
or negative results.
4.
In some cases, sample preparation can be complex and
time-consuming, requiring special techniques and reagents.
5.
Dot blot can only be used to analyze one target at a time,
making it unsuitable for multiplexing applications.
153
Immunoprecipitation
Immunoprecipitation is a laboratory technique used in molecular
biology and biochemistry to isolate specific proteins from a mixture.
This technique involves using an antibody that specifically binds to
the target protein, and using it to pull out the protein from the
mixture. The protein-antibody complex is then precipitated (or
pelleted) using a reagent such as protein A or protein G, and the
resulting precipitate is washed to remove any non-specific
contaminants. The target protein can then be analyzed by
techniques such as gel electrophoresis or mass spectrometry.
Immunoprecipitation is commonly used in the study of proteinprotein interactions, protein localization, and post-translational
modifications.
Principle
The principle of immunoprecipitation is based on the specific
interaction between an antibody and its antigen. In this technique, a
specific antibody is used to bind to a target protein in a mixture of
proteins. The antibody-antigen complex is then precipitated, or
separated, from the other proteins in the mixture, allowing for the
purification and analysis of the target protein. This method is
commonly used in molecular biology and biochemistry to isolate and
study specific proteins in complex mixtures.
154
Procedure
1.
Prepare the sample by lysing cells or tissues in a lysis buffer.
The sample should contain the target protein and any
interacting proteins.
2.
Prepare the primary antibody that will be used to specifically
bind to the target protein. It is important to choose an
antibody with high specificity for the target protein.
3.
Mix the lysed sample with the primary antibody and
incubate for 1-2 hours at room temperature.
4.
Add protein A/G beads to the sample-antibody mixture and
incubate for another 1-2 hours.
5.
Wash the beads with washing buffer to remove any
6.
Elute the target protein from the beads by adding elution
unbound proteins. Repeat this step 2-3 times.
buffer. The target protein is now ready for further analysis,
such as Western blotting or mass spectrometry.
7.
Analyze the eluted target protein using the chosen method
of analysis. The goal is to identify any interacting proteins
that were immunoprecipitated with the target protein.
Advantages
1.
Immunoprecipitation is based on the binding between an
antibody and its target antigen, which provides high
specificity in selecting a particular protein from a complex
mixture.
2.
It can detect low-abundance proteins that might not be
3.
It can be used for the study of post-translational
detectable by other methods, such as gel electrophoresis.
modifications,
protein-protein
interactions,
and
the
identification of novel proteins.
155
4.
It can be used in various research areas, including cellular
biology, biochemistry, and proteomics.
Disadvantages
1.
Immunoprecipitation is a multi-step process that can take
several
hours
to
complete,
and
can
be
prone
to
contamination and human error.
2.
Antibodies are expensive and must be purchased for each
experiment, making the process cost-prohibitive for some
labs.
3.
Non-specific binding of the antibody to other proteins or
contaminants can occur, leading to false-positive results.
4.
The quality of the antibody can greatly affect the results of
the immunoprecipitation, and antibodies may not be
available for all proteins of interest.
156
Sanger Sequencing
DNA sequencing is the act of finding out the exact order of the
nucleotide bases (As, Ts, Cs, and Gs) in a DNA molecule. This
information is significant in various areas like medical diagnosis,
biotechnology, and forensic biology. There are several techniques for
DNA sequencing, including Maxam-Gilbert sequencing (chemical
degradation
method),
Sanger
sequencing
(dideoxy
chain-
termination method), and high-throughput sequencing technologies.
Among these methods, Sanger sequencing is the most commonly
used and it was developed by Sanger and his team in 1975 due to its
simplicity and reliability.
The Sanger sequencing method employs the cycle sequencing
technique, where dideoxynucleosides are marked with different
fluorescent dyes, allowing all four reactions to occur in the same
tube and be separated in a single lane on the gel. When the labeled
DNA fragments pass through the bottom of the gel, a laser reader
detects the fluorescence of each fragment (blue, green, red, or
yellow) and compiles the data into an image.
The Sanger method is based on the mechanism of DNA synthesis by
DNA polymerases and requires the synthesis of a complementary
DNA strand to the strand being analyzed. This process uses ddNTPs
tagged with fluorescence dye (each nucleotide with a different
color). The identity of the added deoxynucleotide is determined by
its complementarity through base pairing with a base in the
template strand.
157
In the Sanger sequencing reaction, nucleotide analogs called
dideoxynucleoside triphosphates (ddNTPs) interrupt the DNA
synthesis as they lack the 3'-hydroxyl group required for the next
step. For instance, the addition of ddCTP to an otherwise normal
reaction system causes some of the synthesized strands to end
prematurely at the position where dC would normally be added,
opposite a template dG. This results in different colored DNA
fragments, which can be separated by size in an electrophoretic gel
in a capillary tube. All fragments of a given length move together in
a single band through the capillary gel and the color associated with
each band is detected with a laser beam. The DNA sequence is read
by identifying the color sequences in the bands as they pass the
detector, with the amount of fluorescence in each band being
represented as a peak in the computer output.
Procedure
Sanger's Method is a classic DNA sequencing method which involves
the following steps:
1.
A single-stranded DNA template is prepared for sequencing.
This
template
is
usually
derived
from
plasmids,
bacteriophages, or genomic DNA. The template can be
amplified using PCR (Polymerase Chain Reaction) or other
methods.
2.
A short complementary primer, about 20-24 nucleotides in
length, is annealed to the template. The primer is designed
to anneal at the 5’ end of the sequence of interest.
3.
A modified form of DNA polymerase (such as Taq
polymerase) is used to extend the primer in the presence of
four different dNTPs (deoxynucleoside triphosphates). The
dNTPs contain one of the four bases (A, C, G, T), each
labeled with a different fluorescent dye.
158
4.
As the DNA polymerase extends the primer, it encounters a
modified dNTP (ddNTP), which terminates the extension
reaction. This creates a series of fragments of different
lengths, each with a fluorescent label at the end of the
sequence.
5.
The labeled fragments are separated by electrophoresis on a
gel or capillary. The gel or capillary is run at a high voltage,
causing the fragments to separate based on their size.
6.
The fluorescent labels are detected and the resulting data is
analyzed to determine the sequence of the template DNA.
The sequence is determined by analyzing the relative
positions of the fluorescent labels on the gel or capillary.
7.
The individual sequences obtained from the different
reactions are assembled to form the final DNA sequence.
This assembly is done using specialized software that
matches the overlapping regions between the sequences to
determine the final DNA sequence.
Alternate method
1.
Amplify the specific region of DNA using PCR (polymerase
chain reaction). This will result in multiple copies of the
desired DNA fragment.
2.
Purify the PCR product mixture by removing any unwanted
primers and dNTPs (deoxynucleoside triphosphates) to
obtain a pure sample of the DNA fragment.
3.
Perform the sequencing reaction, which uses special dyes to
label the different DNA bases (A, C, G, and T). This will
determine the sequence of the DNA fragment.
4.
Clean up the product after the sequencing reaction to
remove any excess dye terminators and unused primer. This
is done using an ethanol precipitation protocol to purify the
sample.
159
5.
Separate the labeled DNA bases and identify them using
capillary electrophoresis. The resulting data is then analyzed
to produce a final DNA sequence.
6.
Analyze the data obtained from capillary electrophoresis to
produce a complete and accurate DNA sequence. This
sequence will provide valuable information about the DNA
fragment being analyzed.
Advantages
Sanger sequencing has a wide range of applications, including the
detection of single nucleotide polymorphisms (SNPs), single-strand
conformation polymorphism (SSCP), and mutations. It is a reliable
and efficient method for DNA sequencing, making it a popular
choice for many genetic research projects.
160
Maxam-Gilbert Sequencing
DNA sequencing is the process of determining the precise order of
nucleotides within a DNA molecule. The Maxam-Gilbert method, also
known as chemical degradation sequencing, is a chemical-based
DNA sequencing method that was first introduced in 1977. It is
based on the selective chemical modification of DNA strands and the
subsequent analysis of the modified strands.
The Maxam-Gilbert method uses chemicals to selectively break DNA
at specific locations, depending on the type of chemical used. These
modifications generate specific fragments of DNA that can be
separated and identified through electrophoresis. Electrophoresis is
a method that uses an electric field to separate DNA fragments
based on size.
The first step in the Maxam-Gilbert method is to choose a DNA
sample that needs to be sequenced. The sample is then divided into
four aliquots, each of which is treated with a different chemical. The
chemicals used in the Maxam-Gilbert method are hydroxylamine,
which cleaves the DNA at purine bases, and a combination of
chemicals called G, A, and C, which cleave the DNA at guanine,
adenine, and cytosine bases, respectively.
After treatment with the chemicals, the DNA fragments are
separated by electrophoresis and the resulting bands are visualized
using a method such as autoradiography. Autoradiography uses Xray film to detect radioactive isotopes that have been incorporated
into the DNA samples during treatment.
161
Procedure
The Maxam-Gilbert method is a chemical cleavage method for DNA
sequencing. The steps for DNA sequencing through Maxam-Gilbert's
method are as follows:
1.
Obtain a sample of DNA from a suitable source such as
bacteria or human cells. The DNA should be pure, free from
contaminants and suitable for sequencing.
2.
Cut the DNA sample into smaller fragments using restriction
enzymes. These enzymes recognize specific DNA sequences
and cut the DNA at these sites.
3.
Heat the DNA sample to separate the double-stranded DNA
4.
Add specific chemicals to modify the DNA, such as ethidium
into single strands.
bromide, for visualization of the fragments under UV light.
5.
Label the DNA fragments with radioactive phosphorus or
phosphorus-32
(32P)
to
make
them
visible
under
autoradiography.
6.
Load the radiolabeled DNA fragments onto a polyacrylamide
gel. Apply an electric field to the gel, causing the DNA
fragments to move towards the positive electrode. The
smaller fragments move faster and separate from the larger
fragments.
7.
Treat the gel with chemicals, such as hydrazine or nitrous
acid, to cleave the DNA at specific points along the strand.
8.
Expose the gel to X-ray film to visualize the DNA fragments.
The radioactive 32P labels on the DNA produce an image on
the film, which shows the position of each DNA fragment on
the gel.
162
9.
Analyze the autoradiogram to determine the sequence of
the DNA fragments. The position of each fragment on the
gel indicates the sequence of bases in the DNA.
10.
Combine the sequences of the different fragments to obtain
the complete DNA sequence.
Advantages and Disadvantages
The Maxam-Gilbert method has several advantages over other DNA
sequencing methods. One of the main advantages is that it provides
high-resolution data and can be used to sequence large DNA
fragments. Additionally, the Maxam-Gilbert method is relatively
quick and easy to perform and can be automated for highthroughput sequencing.
However, there are also some limitations of the Maxam-Gilbert
method. The method is not as sensitive as other DNA sequencing
methods, and it may not work as well on samples that contain large
amounts of contaminants. Furthermore, the method requires the use
of hazardous chemicals and radioactive isotopes, making it
potentially dangerous and requiring proper safety measures.
163
Pyrosequencing
Pyrosequencing is a type of DNA sequencing technology that uses
bioluminescence to identify the individual nucleotides (A, C, G, T)
that make up a DNA strand. It is based on the principle of real-time
sequencing by synthesis, where the next nucleotide is incorporated
into the growing strand only after the previous one has been
identified. The result of the reaction is a series of light signals, which
are then converted into a DNA sequence. This technology is highly
sensitive, fast, and scalable, making it useful for a wide range of
applications, including genomic sequencing, epigenetic analysis, and
bacterial identification.
Principle
Pyrosequencing is a fast and efficient DNA sequencing method that
uses bioluminescence to identify and quantify the order of
nucleotides in a target DNA sample. The process involves four main
steps: template preparation, primer annealing, enzyme addition, and
detection of incorporated nucleotides. The DNA sample is first
amplified through PCR and mixed with a sequencing primer that
serves as the starting point for the sequencing reaction. The addition
of a sequencing enzyme triggers bioluminescent reactions for each
incorporated nucleotide, producing light that is detected and
processed to determine the DNA sequence. This technology is
unique in its ability to directly detect the incorporation of
nucleotides into a growing DNA strand.
164
Procedure
Pyrosequencing is a method of sequencing DNA that involves the
sequential release of individual nucleotides. The following are the
steps involved in performing pyrosequencing:
1.
The first step is to prepare the DNA sample for sequencing.
This is done by isolating the DNA from the sample, such as
blood or tissue, and then purifying it.
2.
The next step is to amplify the target DNA sequence using
polymerase chain reaction (PCR). This increases the amount
of DNA available for sequencing.
3.
The amplified DNA is then mixed with a primer that is
complementary to the target sequence. This primer allows
the DNA to be attached to a solid support, such as a bead or
a well.
4.
The next step is to add a sequencing enzyme, such as
luciferase or ATP sulfurylase, to the mixture. This enzyme is
responsible for releasing nucleotides in response to the
incorporation of each new base into the growing DNA
strand.
5.
As the DNA strand grows, the sequencing enzyme releases
individual nucleotides, which are detected and quantified by
a luminescence-based assay.
6.
The release of nucleotides is detected by a luminometer,
which measures the amount of light emitted by each
nucleotide. This data is used to determine the sequence of
the DNA.
7.
The data from the luminometer is then analyzed to
determine the sequence of the DNA. This can be done using
computer algorithms that match the data with a reference
genome.
165
8.
Finally, the results of the sequencing analysis are interpreted
to determine the identity of the DNA sample and any
mutations or variations that may be present.
These are the steps involved in performing pyrosequencing. It is a
fast and efficient method of sequencing DNA, and it is widely used in
a variety of applications, including medical genetics and microbial
genomics.
Advantages
1.
Pyrosequencing can sequence hundreds of thousands of
DNA molecules simultaneously, making it a high-throughput
method.
2.
The results of Pyrosequencing are generated in real-time,
3.
It can be used for a wide range of applications, including
which makes it a fast method of sequencing.
single
nucleotide
polymorphism
(SNP)
detection,
transcriptomics, metagenomics, and others.
4.
It is a highly sensitive method, capable of detecting small
amounts of DNA.
5.
Pyrosequencing is relatively less expensive compared to
other sequencing methods, making it an attractive option for
large-scale sequencing projects.
Disadvantages
1.
The maximum read length of pyrosequencing is around 400450 base pairs, which is shorter compared to other
sequencing methods.
2.
The accuracy of pyrosequencing is not as high as other
sequencing methods, making it unsuitable for certain
applications.
166
3.
Pyrosequencing does not have the ability to detect complex
genomic structures, such as insertions, deletions, or
inversions.
4.
This method requires large amounts of starting material,
which can limit its use in certain applications.
5.
The equipment used in pyrosequencing is complex and
requires technical expertise to operate, which can limit its
use in certain settings.
167
Multiplex DNA Sequencing
Multiplex DNA sequencing is a high-throughput sequencing
technique that allows the simultaneous analysis of multiple DNA
samples in a single run. This method involves the addition of unique
barcodes to each sample, which allows for the identification and
separation of the individual sequences once the DNA has been
sequenced. Multiplex DNA sequencing is commonly used in largescale genomic projects, gene expression analysis, and disease
association studies. It offers significant cost savings compared to
sequencing each sample separately, as well as increased efficiency in
the analysis of large numbers of samples.
Principle
Multiplex DNA sequencing is a method of sequencing multiple DNA
samples simultaneously in a single sequencing run. The principle
behind this technique is to distinguish different DNA samples in the
sequencing process by assigning them a unique identifier or
barcode. This identifier is added to the DNA samples before
sequencing, allowing them to be separated and identified later in the
process. The multiplex sequencing process results in highthroughput and cost-effective sequencing of multiple DNA samples,
making it useful for applications such as large-scale genomic studies
and targeted gene sequencing.
168
Procedure
Multiplex DNA sequencing is a technique used to sequence multiple
DNA samples in a single reaction. Here are the steps to perform
multiplex DNA sequencing:
1.
Collect the DNA samples you wish to sequence and extract
the DNA. You will also need to prepare libraries of the DNA
samples, which involves fragmenting the DNA, ligating
adapters, and amplifying the fragments.
2.
Mix the DNA libraries of different samples in a single
reaction to create a pool.
3.
Check the quality of the DNA pool using gel electrophoresis
to confirm that the DNA fragments are of the correct size
and the concentration is adequate for sequencing.
4.
Hybridize the DNA pool with a sequencing bead array, which
is a bead-based technology that contains millions of
individual beads, each with a specific sequence. This allows
for parallel sequencing of multiple samples in a single run.
5.
Perform a PCR reaction in an oil-based emulsion to amplify
the DNA-sequencing bead complexes. This results in many
copies of the bead-DNA complexes that are ready for
sequencing.
6.
Load
the
bead-DNA
complexes
into
a
sequencing
instrument and run the sequencing reaction. The instrument
will read the DNA sequences and generate data that can be
analyzed.
7.
Analyze the data generated by the sequencing reaction to
determine the DNA sequences. You can use bioinformatics
tools to align the sequences and compare them to reference
sequences.
8.
Interpret the data to identify any mutations or variations in
the DNA sequences. This can provide valuable information
169
for a range of applications, including disease diagnosis and
drug development.
Advantages
1.
Multiplex sequencing allows the simultaneous sequencing of
multiple samples in a single run, making it an efficient and
cost-effective solution for large-scale sequencing projects.
2.
By sequencing multiple samples at once, the chances of
detecting errors in the data are reduced, leading to
improved accuracy of the results.
3.
Multiplex sequencing reduces the costs associated with
sequencing as a single run can analyze multiple samples at
once, reducing the need for multiple sequencing runs.
4.
With multiplex sequencing, a wide range of samples can be
analyzed, including samples from different organisms, tissue
types, and genetic backgrounds.
Disadvantages
1.
Multiplex sequencing requires specialized equipment and
technical skills, which can make it challenging to implement
for some researchers.
2.
When multiple samples are analyzed in a single run, the
signals from different samples can interfere with each other,
affecting the quality of the results.
3.
Because
of
the
multiplexing,
the
sensitivity
of
the
sequencing results may be reduced, potentially leading to
missed variants or other important information.
4.
Multiplex sequencing involves a complex process that can be
difficult to manage and troubleshoot, leading to potential
errors in the results.
170
Automated Sequencing
Automated sequencing is a process of determining the sequence of
nucleotides in a DNA sample using automated and computerized
methods. The process involves breaking the DNA sample into
smaller fragments, preparing the fragments for sequencing, running
the sequencing reactions, and analyzing the data to determine the
DNA sequence. Automated sequencing has revolutionized the field
of genomics by making it faster, more efficient, and cost-effective to
sequence large amounts of DNA.
Principle
The principle of automated sequencing is based on the use of
computer-controlled instruments and laboratory methods to analyze
the sequence of nucleotides in a DNA molecule. The process involves
breaking down the DNA into smaller fragments, determining the
sequence of the individual fragments, and then using software
algorithms to reassemble the sequence into a complete genome.
The automated sequencing process can be performed using various
technologies, such as Sanger sequencing, next-generation or highthroughput sequencing, depending on the application and the
desired accuracy and speed of the results. The automated
sequencing process has revolutionized the field of genetics and
molecular biology, enabling researchers to quickly and efficiently
analyze DNA sequences for various applications, such as genetic
diagnosis, genetic engineering, and drug discovery.
Procedure
171
1.
Obtain the DNA sample and extract it to purify and
concentrate the DNA.
2.
Cut the purified DNA into small fragments, usually ranging
from 100-600 base pairs.
3.
Attach primers to the DNA fragments. Primers are short
pieces of complementary DNA that help the sequencing
reaction start at a specific point.
4.
Mix the DNA fragments, primers, and sequencing reagents,
including dNTPs (nucleotides), polymerase, buffer, and
fluorescent dyes.
5.
Perform a series of thermal cycles to extend the primers and
build a complementary strand of DNA using the dNTPs.
6.
Add a sequencing terminator mixture that stops the
polymerase reaction and incorporates fluorescent dyes into
the newly synthesized strand of DNA.
7.
Load the samples into a sequencing machine and analyze
the images produced by the fluorescent dyes to determine
the sequence of the DNA.
8.
Use computer software to analyze the data and produce a
readout of the DNA sequence.
9.
Check the quality of the sequence, including the accuracy
and completeness, and make any necessary adjustments.
10.
Store the sequence data in a secure, accessible location for
future use.
Advantages
1.
Automated sequencing machines can process hundreds or
thousands of samples in a single run, making it faster than
manual sequencing methods.
2.
Automated
sequencing
machines
are
equipped
with
advanced algorithms and software, which ensure high
accuracy and reproducibility of results.
172
3.
Automated sequencing machines reduce the cost per
sample as they are capable of processing many samples
simultaneously, thus reducing the cost per sample.
4.
Automated sequencing machines have the ability to process
large amounts of samples in parallel, making it ideal for
large-scale sequencing projects.
5.
Automated sequencing machines are user-friendly and
require minimal technical skills, making it accessible to a
wide range of users.
Disadvantages
1.
Automated sequencing machines are expensive to purchase,
maintain and repair.
2.
Automated sequencing machines may be limited in their
ability to process small samples or rare specimens.
3.
Automated sequencing machines are complex machines that
require maintenance and technical support.
4.
Automated sequencing machines generate large amounts of
data that requires specialized software and training to
interpret accurately.
5.
Automated sequencing machines may have limitations in
terms of the types of samples they can process, the range of
DNA sequences they can detect, and the types of genetic
mutations they can identify.
173
Construction of Molecular Maps
Construction of molecular maps is the process of creating visual
representations of the spatial relationships and interactions between
atoms, molecules, and other components of a biological system. This
process can involve using different techniques and tools, such as Xray
crystallography,
nuclear
magnetic
resonance
(NMR)
spectroscopy, electron microscopy, and computer simulations, to
obtain detailed information about the structure and function of
biological macromolecules. The information obtained from these
techniques is then used to create two-dimensional or threedimensional molecular maps, which provide insight into the
molecular interactions and processes taking place within the
biological system. These maps can be used for a variety of purposes,
including the design of new drugs, the understanding of biological
processes, and the improvement of medical treatments.
Principle
The principle of construction of molecular maps is to generate a
visual representation of the molecular structure of a substance and
its spatial distribution. This is achieved through the use of various
techniques such as X-ray crystallography, NMR spectroscopy,
electron microscopy, and computer simulations. The molecular maps
provide information about the location and orientation of atoms,
bonds, and functional groups within a molecule, which can be used
to understand its properties, interactions, and reactions with other
molecules.
174
Procedure
1.
Choose a data set - select a set of genetic or molecular
markers.
2.
Determine the genetic variations in each sample using PCR
or sequencing.
3.
Calculate pairwise distances or similarities between markers
and visualize them as a dendrogram or matrix.
4.
Use specialized software to create the molecular map.
5.
Choose highly informative markers based on frequency of
occurrence or ability to distinguish between genotypes.
6.
Add additional markers or re-analyze the data to improve
resolution and identify errors.
7.
Compare the molecular map to other reference datasets to
ensure accuracy and reliability.
Alternative method
1.
Start by identifying the target molecule for which the
molecular map needs to be constructed.
2.
Collect all available information on the target molecule
including its chemical structure, physical properties, and
behavior under different conditions.
3.
Use computer software or tools such as ChemDraw or
ACD/Chem-Sketch to draw the chemical structure of the
target molecule.
4.
Identify the functional groups present in the molecule and
mark them on the chemical structure.
5.
Determine the bond connectivity between different atoms in
the molecule and mark them on the structure.
6.
Identify the stereochemistry of the molecule, if applicable,
and mark it on the structure.
175
7.
Using the marked-up structure, create a molecular map that
clearly
shows
the
different
functional
groups,
bond
connectivity, and stereochemistry of the molecule.
8.
Use appropriate software or tools such as MarvinSketch or
ChemBioDraw to label and annotate the molecular map,
including the chemical name, molecular weight, and any
relevant physical properties.
9.
Validate the accuracy of the molecular map by crosschecking it with the original chemical structure and any
available literature.
10.
Save the molecular map in a suitable file format for further
analysis or use in research.
Advantages
1.
It provides a detailed and accurate view of the molecular
structure of a particular organism or system.
2.
It provides a tool for identifying and studying diseasecausing genes and mutations.
3.
It provides a means to compare the genomes of different
organisms, leading to a better understanding of evolution
and genetic diversity.
Disadvantages
1.
May be subject to errors and inaccuracies due to limitations
in sequencing technology or data analysis methods.
2.
The interpretation of molecular map data can be complex
and may require advanced computational skills.
176
Restriction Mapping
Restriction mapping is a technique used to create a physical map of
a DNA molecule by cutting the DNA at specific locations using
restriction enzymes. The resulting fragments are separated by size
using gel electrophoresis and the pattern of fragments is used to
determine the location and distance between the restriction enzyme
cutting sites. Restriction mapping is used in molecular biology
research to study the organization and structure of DNA, to identify
mutations or genetic variations, and to construct recombinant DNA
molecules.
Principle
The principle of Restriction Mapping is to identify the location of
specific DNA sequences within a DNA molecule by using restriction
enzymes to cleave the molecule at specific recognition sites. The
resulting fragments are then separated by gel electrophoresis,
allowing the size and location of the fragments to be determined.
This technique can be used to create a map of the DNA molecule,
which can aid in gene sequencing and identification.
Procedure
1.
Collect the DNA sample to be analyzed and obtain the
restriction enzymes required for the mapping process.
2.
Digest the DNA sample using the restriction enzymes. This
can be done by adding the enzymes to the sample and
incubating at the appropriate temperature and time for the
enzymes to cut the DNA at their specific recognition sites.
177
3.
Separate the resulting DNA fragments by size using gel
electrophoresis. Load the digested DNA onto an agarose gel
and run an electric current through it to separate the
fragments according to size.
4.
Stain the gel with a DNA-binding dye, such as ethidium
bromide, to visualize the DNA fragments.
5.
Measure the size of the DNA fragments using a DNA ladder
as a reference. A DNA ladder is a mixture of fragments of
known size used to calibrate the size of the fragments in the
sample.
6.
Use the fragment sizes to create a restriction map of the
DNA. This can be done by comparing the fragment sizes to
the expected sizes based on the known recognition sites for
the restriction enzymes used.
7.
Verify the restriction map by repeating the digestion and gel
electrophoresis steps and comparing the resulting fragment
sizes to the predicted sizes on the map.
Advantages
1.
Restriction mapping provides a precise and accurate map of
the DNA sequence, allowing for identification of specific
genetic sequences.
2.
Restriction enzymes can identify and cut specific sequences,
allowing for the identification of specific genes or mutations.
3.
Restriction mapping allows for the analysis of DNA
sequences, such as the identification of deletions or
insertions.
4.
Restriction mapping can be used in genetic mapping, which
is important in understanding hereditary diseases.
178
Disadvantages
1.
Restriction mapping can be a time-consuming process,
requiring significant amounts of time and resources.
2.
The process of restriction mapping can be expensive, as it
requires specialized equipment and reagents.
3.
Restriction mapping can be complex, and the interpretation
of the results may require significant expertise.
4.
Restriction mapping is limited to specific sequences
recognized by restriction enzymes, which may not be
sufficient for certain genetic analyses.
179
Molecular Markers
Molecular markers are specific DNA sequences or variations that can
be used to identify and differentiate individuals or populations of
organisms. They are important tools in genetic research, plant and
animal breeding, and forensic science.
Restriction Fragment Length Polymorphism
Restriction Fragment Length Polymorphism (RFLP) is a technique
that involves digesting DNA with restriction enzymes and separating
the resulting fragments by gel electrophoresis. The resulting pattern
of fragments can be used to identify differences between individuals
or populations.
Principle
Restriction Fragment Length Polymorphism (RFLP) is a molecular
biology technique used to detect variations in DNA sequences
between different individuals. The principle of RFLP is based on the
fact that DNA sequences vary from person to person, and that these
variations can be detected by analyzing the patterns of DNA
fragments generated by restriction enzymes.
Restriction enzymes are enzymes that cut DNA at specific sites,
creating fragments of different sizes. By digesting DNA samples from
different individuals with the same restriction enzyme, researchers
can compare the resulting fragments and identify differences in the
DNA sequences between the samples.
180
These differences can be visualized using gel electrophoresis, a
technique that separates DNA fragments based on their size. The
resulting pattern of DNA fragments, or DNA fingerprint, can be used
to identify individuals, establish relationships between individuals, or
detect genetic mutations associated with diseases.
Overall, the principle of RFLP is based on the idea that variations in
DNA sequences can be detected and analyzed by digesting DNA
samples with restriction enzymes and visualizing the resulting
fragments using gel electrophoresis.
Stock
DNA sample
EcoRI restriction endonuclease enzyme
Agarose, ethidium bromide
Electrophoresis apparatus
Gel documentation system
Procedure
1.
Start by obtaining a DNA sample from the organism you
wish to analyze. This can be done by collecting a tissue
sample or blood sample from the organism.
2.
Extract the DNA from the sample using a DNA extraction kit.
Follow the manufacturer's instructions for the kit.
3.
Take the extracted DNA and cut it using a restriction
enzyme. This enzyme recognizes a specific sequence of DNA
and cuts it at a specific site. This produces fragments of
varying sizes.
4.
Separate the DNA fragments using gel electrophoresis. Place
the DNA fragments in a gel and run an electric current
through the gel. This separates the fragments by size.
181
5.
Transfer the separated DNA fragments to a membrane, such
as a nitrocellulose or nylon membrane. This is done using a
technique called Southern blotting.
6.
Hybridize the membrane with a labeled probe that will bind
to a specific DNA sequence. The probe will bind to the DNA
fragments on the membrane that contain the sequence it
recognizes.
7.
Use an imaging system to visualize the labeled probe and
the DNA fragments it has bound to. This will produce a
pattern of bands on the membrane.
8.
Analyze the band pattern to determine the genotype of the
organism. This can be done by comparing the band pattern
to known patterns for the organism or by using computer
software to analyze the pattern.
Random Amplified Polymorphic DNA
Random Amplified Polymorphic DNA (RAPD) is a PCR-based
technique that amplifies random fragments of DNA using short,
random primers. The resulting pattern of amplified fragments can be
used to identify differences between individuals or populations.
Principle
RAPD is a PCR-based technique that uses short, arbitrary primers to
amplify DNA fragments that contain polymorphic regions. The
principle of RAPD is based on the fact that the primers used in the
reaction are random and do not target specific regions of the DNA.
This results in the amplification of a large number of fragments that
are specific to the genomic DNA of the sample being tested. The
amplified fragments are then separated by gel electrophoresis and
visualized to identify polymorphic regions in the DNA. RAPD is a
useful technique for identifying genetic variability within populations,
182
for fingerprinting organisms, and for studying genetic relationships
between different organisms.
Procedure
Here is the step-wise procedure to perform RAPD:
1.
Collect DNA samples from the organisms you want to study.
Make sure to extract high-quality DNA using a suitable
extraction method.
2.
Prepare the PCR reaction mixture by adding the template
DNA, primers, Taq polymerase, and other necessary
components in the right proportions. Mix the reaction
mixture well.
3.
Set up the PCR amplification by placing the reaction mixture
in a thermal cycler. Run the PCR reaction for the specified
number of cycles, with appropriate temperature and time
conditions.
4.
Once the PCR amplification is completed, visualize the
amplified DNA fragments by running the reaction products
on an agarose gel. Stain the gel with an appropriate dye and
visualize the bands under UV light.
5.
Analyze the RAPD banding patterns by comparing the
banding patterns of different DNA samples. The presence or
absence of particular bands can be used to identify genetic
variations among the studied organisms.
6.
Interpret the results of RAPD analysis based on the banding
patterns obtained. Use appropriate statistical tools to
analyze and compare the results.
7.
Record the findings of the RAPD analysis and document
them in a clear and concise manner.
8.
Draw conclusions based on the RAPD analysis results and
their significance for the research question or objective.
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9.
Repeat the RAPD experiment if necessary to confirm the
results or to address any additional research questions.
Amplified Fragment Length Polymorphism
Amplified Fragment Length Polymorphism (AFLP) is a PCR-based
technique that combines the use of restriction enzymes and selective
amplification of fragments using specific primers. The resulting
pattern of amplified fragments can be used to identify differences
between individuals or populations.
Principle
Amplified Fragment Length Polymorphism (AFLP) is a molecular
genetic technique that is used to study genetic variation in different
organisms. The principle of AFLP involves the use of selective PCR
amplification of genomic DNA fragments that are flanked by
restriction sites, followed by the separation of these fragments by
electrophoresis and detection by autoradiography or fluorescence.
AFLP allows for the detection of thousands of DNA polymorphisms
at a time, providing a high-resolution genetic fingerprint of the
individual or population under study. This technique is widely used in
population genetics, evolutionary biology, plant breeding, and
molecular systematics.
Procedure
1.
Start by isolating DNA from the tissue or organism of
2.
Cut the DNA using restriction enzymes that recognize
interest using a suitable protocol.
specific sequences, which will generate fragments of varying
lengths.
3.
Add specific adapter sequences to the ends of the DNA
fragments using ligases.
184
4.
Amplify the DNA fragments using PCR with adapter-specific
primers.
5.
Use a set of selective PCR primers to amplify a subset of
fragments that contain the restriction enzyme recognition
site and the adapter sequence.
6.
Separate the amplified fragments using gel electrophoresis,
and visualize them using a suitable method, such as staining
or autoradiography.
7.
Analyze the resulting fragment patterns to determine
genetic variation among the samples. This can be done by
creating a dendrogram or performing statistical analysis to
cluster similar samples together.
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DNA Ship and Microarrays
DNA chip and microarrays are high-throughput methods that
involve immobilizing thousands of DNA fragments on a solid surface
and hybridizing them with labeled probes. The resulting pattern of
hybridization can be used to identify differences in gene expression
or genetic variation between individuals or populations. These
techniques can be used for genotyping, gene expression profiling,
and other applications in genetics and biotechnology.
Principle
The principle of DNA chip, also known as microarray technology, is
based on the ability to simultaneously analyze the expression or
sequence of thousands of genes or DNA sequences in a single
experiment. A DNA chip is a small glass slide or silicon chip on which
thousands of microscopic spots or probes have been immobilized,
each of which contains a unique DNA sequence. The samples
containing DNA fragments are labeled with fluorescent dyes and
hybridized to the probes on the chip. The intensity of the
fluorescence signal is then measured, allowing researchers to identify
which genes are expressed or which DNA sequences are present in
the sample. This technology is widely used in genomics research,
diagnosis, and personalized medicine.
Procedure
Performing DNA and Microarray Procedure
1.
The first step in DNA and microarray procedure is to collect
the sample. It could be blood, tissue, or any other material
186
that contains DNA. Make sure to collect a sufficient amount
of sample for the experiment.
2.
After collecting the sample, the next step is to isolate DNA
from it. You can use a DNA isolation kit for this purpose.
Follow the manufacturer's instructions to extract DNA.
3.
Once you have isolated DNA, you need to measure its
concentration using a spectrophotometer. The concentration
of DNA should be within the required range for the
microarray experiment.
4.
The next step is to label the DNA using fluorescent dyes.
This process will enable the microarray scanner to detect the
DNA hybridization on the microarray slide.
5.
The microarray slide contains thousands of DNA probes that
can hybridize with the labeled DNA. Before adding labeled
DNA to the microarray slide, wash the slide with buffer
solution to remove any impurities.
6.
Add the labeled DNA to the microarray slide and incubate it
for several hours. During this time, the labeled DNA will
hybridize with the complementary probes on the slide.
7.
After incubation, scan the microarray slide using a
microarray scanner. The scanner will detect the hybridized
DNA probes and provide you with the data.
8.
After scanning the microarray slide, you will get a data file
that contains information about the hybridization patterns.
Analyze this data using bioinformatics tools to determine
gene expression levels or genetic variations.
9.
Based on the analysis of the microarray data, you can
interpret the results and draw conclusions. This information
can be used to diagnose diseases, study genetic variations,
or develop new drugs.
10.
Finally, include all the relevant data and statistical analyses
to draw precise conclusions.
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Advantages
1.
The DNA chip and microarray technology allows for the
simultaneous analysis of thousands of genes in a single
experiment, making it a highly efficient and cost-effective
method.
2.
DNA chips and microarrays are highly accurate in detecting
gene expression levels, enabling researchers to obtain highly
precise and reliable data.
3.
DNA chips and microarrays allow researchers to perform
experiments much faster than traditional methods, saving
time and resources.
4.
DNA chips and microarrays produce large amounts of data,
allowing researchers to analyze the expression of thousands
of genes in a single experiment.
Disadvantages
1.
DNA chips and microarrays only provide information about
the expression of genes that are included in the array, so
they may not be suitable for studying genes that are not
present on the array.
2.
The cost of DNA chips and microarrays can be high,
especially if custom arrays are required.
3.
Interpreting the large amounts of data produced by DNA
chips and microarrays can be challenging, and requires
advanced data analysis skills.
4.
The technology used in DNA chips and microarrays can be
complex, and requires specialized equipment and expertise
to run experiments and interpret data.
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Genomic Library
A genomic library is a collection of DNA fragments that represent
the complete genome of an organism. The fragments are usually
cloned into a suitable vector, such as a bacteriophage or plasmid,
and transformed into bacteria to create multiple copies. The
genomic library serves as a valuable resource for studying the
genetic makeup of an organism and is used in various applications
such as sequencing the genome, identifying disease-causing genes,
and developing new medicines.
Principle
The principle of Genomic Library is to create a collection of cloned
DNA fragments representing the entire genome of an organism.
These cloned fragments can then be used to study the genome and
identify specific genes, their functions, and their interactions. This is
done by randomly fragmenting the genome, cloning each fragment
into a vector, and then transforming the vectors into host cells. The
resulting library of cloned fragments can be screened to isolate and
study specific genes of interest. The principle of genomic library is to
provide a comprehensive representation of the genome for
functional analysis and is a valuable tool for genetic research.
Procedure
1.
First of all, isolate high-quality and high molecular weight
genomic DNA from the sample. This can be done using
commercial kits or by using a protocol such as phenolchloroform extraction or organic extraction.
189
2.
Then, cut the isolated genomic DNA into smaller fragments
using restriction enzymes. The choice of restriction enzymes
will depend on the size and complexity of the genome and
the desired size of the library.
3.
The cut DNA fragments are then separated by size using gel
electrophoresis. The target fragment size for the library
should be within the range of the insert size of the cloning
vector.
4.
The selected fragments are then ligated to a suitable cloning
vector. The cloning vector should have the necessary
elements for replication in bacteria and contain a selection
marker.
5.
The ligated vectors are then transformed into a host
organism, such as E. coli. The host organism takes up the
vector and replicates the DNA fragments, creating a large
number of copies.
6.
The transformed host organisms are then grown on selective
media to identify those that contain the vector with the DNA
fragment of interest. These clones are then screened using
techniques such as hybridization, PCR, or restriction analysis.
7.
The DNA fragment of interest is then sequenced using
techniques such as Sanger sequencing, next-generation
sequencing
(NGS),
or
genome-wide
sequencing.
The
sequencing process determines the order of the DNA base
pairs in the fragment, providing information on the gene
sequence.
8.
The data generated from the sequencing process is then
analyzed to identify the gene and its function. This
information can be used to better understand the biology of
the organism and its role in various processes.
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Advantages
1.
A genomic library provides a comprehensive representation
of an organism's genetic information.
2.
It allows the identification of all genes present in the
genome of an organism.
3.
It facilitates the study of the functions and interactions of
individual genes.
4.
It can be used for genetic screening and disease diagnosis.
5.
It provides a useful tool for genetic engineering and
biotechnology.
6.
It can be used to study gene expression patterns, gene
regulation, and the effects of mutations.
Disadvantages
1.
The process of creating a genomic library is complex, timeconsuming, and expensive.
2.
The genomic library may not be a complete representation
of the genome, as some regions may not be accessible for
cloning.
3.
The genome sequence may contain a large number of
repetitive or redundant elements, making it difficult to
distinguish between similar sequences.
4.
The genomic library may not accurately reflect the functional
state of the genome, as the DNA sequences may not be in
their normal, active configuration.
5.
There is a risk of contamination or mislabeling of the
6.
The genomic library may not be suitable for studying specific
genomic library, leading to incorrect results.
regions of interest, such as those involved in complex
biological processes.
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cDNA Library
cDNA library is a collection of complementary DNA (cDNA)
molecules that represent the complete set of expressed genes in a
particular cell or tissue. The cDNA is synthesized from messenger
RNA (mRNA) molecules using the reverse transcription process and
is then cloned into a vector for storage and replication in a host
organism. The cDNA library can be used to study gene expression
patterns, to identify new genes, or to clone specific genes for
functional analysis. The library serves as a source of material for
molecular biology experiments and is a valuable resource for
understanding
the
molecular
mechanisms
underlying
cellular
processes.
Principle
The principle of cDNA library is based on the reverse transcription of
messenger RNA (mRNA) into complementary DNA (cDNA). The
cDNA library is a collection of cDNA molecules that represent the
complete set of expressed genes in a particular cell, tissue, or
organism.
The process of creating a cDNA library involves the following steps:
1.
Isolation of mRNA: Total mRNA is extracted from the target
tissue or cell.
2.
Reverse transcription: The mRNA is reverse transcribed into
cDNA
using
reverse
transcriptase,
an
enzyme
that
synthesizes DNA from RNA.
192
3.
Cloning: The cDNA molecules are then cloned into a suitable
vector, such as a plasmid or a phage, to generate a library of
cDNA clones.
4.
Screening: The library is screened for specific cDNA clones
representing the genes of interest.
The cDNA library can be used for various applications, including
gene expression analysis, functional genomics, and the identification
of novel genes.
Procedure
Step 1: Isolation of Total RNA
1.
Obtain a sample of the tissue you wish to study (e.g. brain,
liver, muscle).
2.
Grind the tissue to a fine powder in liquid nitrogen.
3.
Extract total RNA using a kit such as Trizol or RNeasy.
Step 2: Synthesis of First-Strand cDNA
1.
Add a reverse transcriptase enzyme, along with random
primers, to the total RNA sample.
2.
Incubate the mixture at 37°C for 60 minutes.
3.
Stop the reaction by heating the mixture to 70°C for 10
minutes.
Step 3: Synthesis of Second-Strand cDNA
1.
Add a DNA polymerase, ligase, and buffer to the first-strand
cDNA.
2.
Incubate the mixture at 16°C for 2 hours.
3.
Stop the reaction by heating the mixture to 70°C for 10
minutes.
193
Step 4: Preparation of cDNA Library
1.
Purify the cDNA using a column-based purification kit.
2.
Add restriction enzymes to the cDNA to cut the DNA into
fragments of desired size.
3.
Ligate the fragments to vectors (e.g. plasmids) using a DNA
ligase.
4.
Transform the ligation mixture into bacteria such as E. coli.
5.
Screen the bacteria for successful transformation and pick
individual colonies for further analysis.
Step 5: Validation of cDNA Library
1.
Isolate plasmid DNA from the transformed bacteria.
2.
Amplify a portion of the cDNA using PCR.
3.
Sequence the amplified cDNA to confirm the presence of
target genes.
4.
Analyze the cDNA library using techniques such as Northern
blotting or qPCR to verify the quality of the library.
Advantages
1.
cDNA libraries are used in sequencing as they provide a
high-quality representation of the mRNA in the sample.
2.
cDNA libraries are constructed from a mixture of mRNA
species and provide comprehensive coverage of the
transcriptome.
3.
cDNA libraries are relatively easy to handle and can be
processed with existing lab equipment and procedures.
4.
cDNA libraries provide consistent and reliable data and are
less prone to technical errors than other types of libraries.
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Disadvantages
1.
cDNA libraries do not always provide a representative
sample of the transcriptome, as only the most abundant
transcripts may be included.
2.
cDNA libraries may miss rare transcripts, particularly those
that are poorly represented in the mRNA pool.
3.
The technical limitations of cDNA library construction can
impact the quality and representativeness of the data
obtained.
4.
The construction of cDNA libraries can be expensive and
time-consuming, requiring specialized equipment and skills.
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DNA Fingerprinting
DNA fingerprinting, also known as DNA profiling, is a laboratory
technique
used
to
identify
an
individual's
unique
genetic
information. This information is collected from a sample of an
individual's DNA, such as blood, saliva, or hair, and compared to
other DNA samples to determine the likelihood of a genetic
relationship between two individuals. DNA fingerprinting is used in
many applications, including forensic investigations, parentage
testing, and genealogy research.
Principle
DNA fingerprinting is based on the principle of genetic variation.
Every individual has a unique DNA sequence, except for identical
twins, which allows them to be identified by their specific genetic
code. This technique uses restriction fragment length polymorphism
(RFLP) to compare the DNA samples from different sources. In RFLP,
restriction enzymes are used to cut the DNA into fragments and then
the fragments are separated by gel electrophoresis. The pattern of
the fragments is unique for each individual and can be used to
identify them. The principle of DNA fingerprinting is used in forensic
science, paternity testing, and other applications where the
identification of individuals is required.
Procedure
DNA fingerprinting, also known as DNA profiling, is a laboratory
technique used to identify individuals based on their unique DNA
patterns. Here is a step-by-step procedure for DNA fingerprinting:
196
1.
Obtain a DNA sample from sources such as blood, saliva,
semen, hair follicles, or skin cells.
2.
Separate DNA from cellular components, such as proteins
and lipids, to obtain pure DNA.
3.
Use polymerase chain reaction (PCR) to create many copies
of a specific target region in the DNA sample.
4.
Cut the amplified DNA into fragments using restriction
enzymes, which recognize specific sequences in the DNA.
5.
Separate the fragments by size using an electric field in a gel
matrix. Smaller fragments will move faster and end up
farther from the point of origin.
6.
Transfer the separated fragments to a nitrocellulose or nylon
membrane. Fix the transferred DNA to the membrane using
baking or ultraviolet light.
7.
Probe the transferred DNA with a labeled complementary
DNA probe to bind specifically to the target DNA.
8.
Expose the membrane to X-ray film to detect the radioactive
label on the probe. This will create a unique fingerprint for
the individual.
9.
Compare the DNA fingerprint with other fingerprints to
determine if they are from the same individual.
Advantages
1.
DNA fingerprinting provides a highly accurate method of
identifying individuals based on their unique genetic
material. This helps in resolving disputes related to identity,
such as in criminal investigations and parentage testing.
2.
DNA evidence is considered very reliable in court and is
often used to determine guilt or innocence. It is difficult to
tamper with and is considered as a robust tool in criminal
investigations.
197
3.
DNA fingerprinting has been instrumental in solving many
crimes, including serial killings, sexual assaults, and other
violent crimes. It can also be used to eliminate suspects,
leaving behind only those who are most likely to be
responsible.
4.
The process of DNA fingerprinting has become much faster
and more efficient over the years, making it easier for
investigators to identify criminals quickly.
Disadvantages
1.
DNA fingerprinting is a relatively expensive process,
especially when compared to other forms of evidence. This
can make it difficult for law enforcement agencies and other
organizations to use it in all cases.
2.
DNA fingerprinting requires special equipment and highly
trained professionals. Technical errors or incorrect analysis
can
lead
to
false results and mistakes in criminal
investigations.
3.
The use of DNA fingerprinting raises serious privacy
concerns
as
it
involves
collecting
sensitive
personal
information. There are also concerns about the misuse of
DNA samples and the possibility of genetic discrimination.
4.
DNA fingerprinting is not always useful in all cases. For
example, it may not provide results if the sample size is too
small, or if it is degraded due to age or environmental
factors.
198
Genetic Selection and Screening
Method
Use of chromogenic substrates
Genetic selection and screening through the use of chromogenic
substrates is a technique that is commonly used in molecular biology
to identify and select specific genes or genetic traits. Chromogenic
substrates are small molecules that can be used to detect the
presence of a specific enzyme or protein. In genetic selection and
screening, these substrates are used to identify cells or organisms
that express a desired gene or trait.
For example, a researcher may want to identify cells that produce a
specific enzyme that is involved in a metabolic pathway. They can
use a chromogenic substrate that is cleaved by the enzyme,
producing a colored product. Cells that produce the desired enzyme
will show a color change, allowing the researcher to select and
isolate these cells.
This technique can also be used to screen for genetic mutations or
variations. By using chromogenic substrates that are specific for
different genetic variants, researchers can identify individuals or
organisms with particular genetic traits or mutations.
Overall, genetic selection and screening through the use of
chromogenic substrates is a powerful tool in molecular biology that
allows for the identification and selection of specific genes and traits.
199
Principle
The principle of genetic selection and screening by the use of
chromogenic substrates involves the use of specific substrates that
are designed to interact with enzymes produced by genetically
modified organisms (GMOs). These substrates are often color-coded,
so that the presence of a particular color indicates the presence of
the desired enzyme or genetic trait.
In this process, scientists modify the DNA of an organism to produce
a specific enzyme that can be detected by the use of a chromogenic
substrate. They then introduce the modified organism into a culture
or environment that contains the substrate. If the organism produces
the desired enzyme, it will interact with the substrate and produce a
color change, allowing researchers to identify and select those
organisms that possess the desired genetic trait.
This process is commonly used in biotechnology to identify and
select organisms that have been genetically modified for specific
purposes, such as producing pharmaceuticals, enzymes, or other
valuable products. It is a powerful tool for genetic engineering and
has revolutionized the field of biotechnology.
Procedure
1.
Prepare a culture of the microorganisms you want to select
or screen. Make sure you grow the culture under the
conditions that favor the expression of the desired
phenotype.
2.
Select a suitable chromogenic substrate that produces a
colored product when acted upon by the enzyme or protein
of interest. Prepare the substrate according to the
manufacturer's instructions.
200
3.
Add the prepared chromogenic substrate to the culture
containing the microorganisms. Make sure you add the
substrate at a concentration that is sufficient to detect the
desired phenotype.
4.
Incubate the culture at the appropriate temperature and for
the necessary amount of time to allow for the desired
enzymatic reaction to occur.
5.
Observe the culture for the presence of colored colonies or
areas. The color development will indicate the presence of
the enzyme or protein of interest, and thus the phenotype
you are selecting for or screening against.
6.
Confirm the presence of the desired phenotype by
performing further tests such as genetic analysis or
biochemical assays.
7.
If necessary, repeat the process with different chromogenic
substrates or under different conditions to further refine the
selection or screening process.
Advantages
1.
Genetic
selection
and
screening
with
chromogenic
substrates allow for a quick and easy analysis of a large
number of samples.
2.
It is a cost-effective method, which reduces the time and
cost of analysis.
3.
Chromogenic substrates are specific and sensitive, which
helps to identify the targeted genes with high accuracy.
4.
It enables the detection of mutations in DNA or protein
sequence that can be difficult to identify using other
methods.
5.
It is a non-destructive method, which can preserve the
sample for further analysis.
201
Disadvantages
1.
Chromogenic substrates have limitations in detecting
mutations that occur outside of the specific target area.
2.
False positives and false negatives can occur due to the
specificity of the chromogenic substrate and the method
used for screening.
3.
The interpretation of results can be subjective, leading to
inconsistencies in the analysis.
4.
Genetic
selection
and
screening
with
chromogenic
substrates can only detect changes in the DNA or protein
sequence, and may not be able to identify epigenetic
changes that influence gene expression.
5.
It may require specialized equipment and expertise, which
can limit the accessibility of the method for some
researchers or institutions.
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Antibiotic Sensitivity Test for
Detection of Recombinants
Principle
To determine the ability of organisms to produce mutants that are
resistant, a gradient plate of a particular antibiotic can be used. This
method involves growing bacteria on a gradient plate, which consists
of two wedge-like layers of media, a layer of plain nutrients, and a
top layer of antibiotic with a nutrient layer.
The antibiotic is added as a top layer to the bottom layer, which
produces an agar gradient of antibiotic concentration from low to
high. Streptomycin is used to create the gradient plate.
E.Coli, which is normally sensitive to streptomycin, will be spread
over the surface of the plates and incubated for 24 to 72 hours.
After inoculation, colonies will appear on the gradients. The colonies
that develop in the high concentration are resistant to the action of
the streptomycin and are considered as streptomycin-resistant
mutants.
For the isolation of antibiotic-resistant growth, the commonly used
antibiotics are Rifampicin, Streptomycin, and Erythromycin.
Stock
NAM
Petri-plates
Glass rod
Sterile test tube
203
Sterile swab
Streptomycin
Procedure
1.
Pour 50 ml of nutrient agar into a sterile petri-plate and
allow the medium to solidify in a standing position by
placing a glass rod under one side.
2.
Once the agar medium is solidified, remove the glass rod
and place the plate in a horizontal position.
3.
Pour nutrient agar with streptomycin (100 gm/ml) solution
into the petri-plate.
4.
Allow the medium to solidify.
5.
Label the low and high antibiotic concentration areas on the
6.
Pipette out 200 ml of 24-hour culture onto the gradient
bottom of the petri-plate.
plate.
7.
Incubate the plate in an inverted position at 37°C for 48-72
hours.
8.
Observe the plate for the appearance of colonies in the area
of
Low
Streptomycin
Concentration
(LSC)
and
High
Streptomycin Concentration (HSC).
Alternative Method
1.
Take a loop and touch the tops of 3-5 colonies of the same
type of organisms from the primary culture plate to prepare
the inoculum.
2.
Transfer the growth to a tube containing saline solution.
3.
Adjust the density of the test suspension to the standard
turbidity by adding either bacteria or sterile saline. Compare
the tube's turbidity with the standard.
4.
Incubate the plates by dipping a swab into the inoculum and
remove excess inoculum.
204
5.
Rotate the plates through an angle of 60° after streaking the
swab over the medium's surface three times.
6.
Pass the swab around the edge of the agar surface.
7.
Close the lid and let the inoculum dry at room temperature
for a few minutes.
8.
Place the antibiotics on the incubated plates using a
template, sterile needle tip, or antibiotic dispenser.
Result
The gradient plate method produced successful results, with lower
growth in the region of the plate without antibiotics compared to
the high antibiotic concentration area.
Notes
Ensure that the slanting position is not disturbed.
Make sure accurate pipetting.
Closely observe the incubated plates.
205
Insertional Inactivation
Genetic selection and screening through insertional inactivation is a
method of identifying genes that are essential for the growth and
survival of an organism. This method involves the insertion of a DNA
sequence, usually a transposon, into the genome of the organism.
This insertion disrupts the function of the gene in which it lands. By
selecting for cells that have lost the ability to grow or survive in a
particular environment, researchers can identify the genes that are
important for that function. This technique is commonly used in the
study of bacteria and other microorganisms, as well as in the genetic
engineering of plants and animals.
Principle
The principle of genetic selection and screening through insertional
inactivation involves the use of genetic engineering techniques to
insert a DNA sequence (usually a marker gene) into a specific
location within the genome of a cell or organism. This DNA
sequence disrupts the normal function of the gene(s) at that
location, resulting in a phenotype that can be selected or screened
for.
For example, in bacterial genetics, a plasmid carrying a marker gene
(such as antibiotic resistance) is introduced into a population of
bacteria. The bacteria that take up the plasmid will have the marker
gene integrated into their genome, disrupting the function of one or
more essential genes. By growing the bacteria on a selective medium
(containing the antibiotic), only those with the marker gene will
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survive and grow. This allows for the selection of bacteria with
specific phenotypes (e.g. antibiotic resistance).
In eukaryotic genetics, similar techniques can be used to create
knockout mice, where specific genes have been disrupted using
marker genes. By breeding these mice and observing the resulting
phenotypes, researchers can study the function of these genes in
development and disease.
Procedure
1.
Start by identifying the gene that you want to inactivate
using insertional mutagenesis. This could be a gene that is
important for a particular biological process or one that is
responsible for a particular trait.
2.
Design a DNA construct that can be used to insert a
selectable marker into the target gene. The construct should
include a selectable marker, such as antibiotic resistance, and
a promoter that can drive expression of the marker.
3.
Introduce the DNA construct into the cells that you want to
mutate. This can be done using a variety of techniques,
including electroporation, lipofection, or viral transduction.
4.
After introducing the DNA construct, select for cells that
have incorporated the selectable marker into the target
gene. This can be done using a selectable antibiotic or other
selectable markers.
5.
Once you have selected for cells with the inserted marker,
screen for cells that exhibit the desired phenotype. This
could be a change in morphology, growth rate, or some
other observable characteristic.
6.
Once you have identified cells with the desired phenotype,
confirm that the mutation is due to the insertional
inactivation of the target gene. This can be done using a
207
variety of techniques, including PCR, sequencing, or
functional assays.
7.
If necessary, repeat the process to identify additional cells
with the desired phenotype or to optimize the mutation.
8.
Finally, characterize the mutant phenotype by studying the
effects of the gene inactivation on the biological process or
trait of interest. This may involve additional experiments,
such as gene expression analysis, biochemical assays, or
animal studies.
Advantages
1.
Insertional inactivation is an efficient method of genetic
screening as it can target a large number of genes at once.
2.
Insertional inactivation can be highly specific in targeting
particular genes, allowing for precise genetic modifications.
3.
The screening process can be relatively quick, taking only a
few days to identify which genes have been disrupted.
4.
This method can be used in a wide range of organisms,
including bacteria, yeast, plants, and animals.
Disadvantages
1.
Insertional inactivation can have off-target effects (in other
genes), which can have unintended consequences.
2.
As much of the genome remains uncharacterized, it is
difficult to know which genes are important to disrupt.
3.
Insertional inactivation may also affect regulatory regions
that control gene expression, leading to unpredictable
effects.
4.
The process of creating and screening a large library of
mutants can be complex and require specialized skills and
equipment.
208
Complementation of Defined Mutations
Genetic selection and screening through complementation of
defined mutations is a method used to identify and study specific
genes and their functions. This technique involves inducing
mutations in a population of cells or organisms, and then selecting
or screening for those with a particular phenotype of interest.
Complementation is a process by which a functional copy of a gene
can restore the phenotype of a mutant with a non-functional copy of
the same gene. By introducing a plasmid or other genetic element
carrying a functional version of the mutated gene into the mutant,
researchers can determine if the phenotype is restored, indicating
that the gene is responsible for the phenotype.
This method is commonly used in genetic research to identify the
functions of unknown genes, study gene interactions, and develop
gene therapies. It is also used in plant and animal breeding to select
for desired traits.
Principle
The principle of genetic selection and screening through complementation of defined mutations involves the use of genetically
modified organisms to identify and study the functions of specific
genes. This approach relies on the ability of cells to compensate for
the loss of a gene function by complementing it with a functional
copy of the gene.
In this method, a mutant strain with a defined mutation in a gene of
interest is crossed with another strain that has a second mutation in
209
the same gene, but at a different site. The resulting progeny are
screened for complementation, which occurs when the two
mutations together restore the wild-type phenotype.
This process enables researchers to identify and study the functions
of specific genes by analyzing the complementation patterns of the
mutant strains. It can also be used to identify potential drug targets,
as
well
as
to
study
genetic
diseases
and
developmental
abnormalities.
Procedure
1.
Pick the set of mutations that need to be screened for
genetic selection. This could include a set of randomly
generated mutations or a specific set of mutations that are
suspected to be linked to a particular phenotype.
2.
Generate a complementation library that contains a set of
plasmids or genomic fragments that express wild-type
copies of the mutated genes. The library should be designed
to cover all the mutations that need to be screened.
3.
Transform the complementation library into a suitable host
strain that is deficient in the specific function being studied.
This is usually achieved by using a host strain that has a
specific deletion or mutation in the gene of interest.
4.
Plate the transformed host strain on a selective media that
does not support growth of the host strain. Only the
transformed host strains that express the wild-type copies of
the mutated genes will be able to grow on the selective
media. This is because the wild-type copies complement the
mutations in the host strain.
5.
Isolate the plasmids that complement the mutation(s) of
interest from the complementation library.
210
6.
Verify the complementation of the isolated plasmids by
retransforming them into the original host strain and
retesting for growth on the selective media.
7.
Characterize the complementing plasmids by sequencing
and mapping them to identify the wild-type genes that
complement the mutated genes.
8.
Repeat the screening process with the remaining mutations
of interest until all the mutations have been screened and
complementing
plasmids
have
been
identified
and
characterized.
9.
Analyze the results to identify the genes that are responsible
for the observed phenotype and gain a better understanding
of the genetic basis of the phenotype.
Advantages
1.
This method allows for very precise selection and screening
of specific mutations. This can help researchers identify the
effects of particular mutations on gene function and disease.
2.
It is very efficient method in terms of selecting the desired
mutation. It requires minimal screening and can quickly
identify the required mutations.
3.
Since this method is based on complementation, it ensures a
high degree of reliability in the screening process.
4.
The cost of genetic selection and screening through complementation is relatively low compared to other methods.
Disadvantages
1.
This method is limited in terms of the mutations it can select
for. It only works for mutations that can be complemented
by a wild-type gene.
2.
Although this method is efficient, it can be time-consuming
to generate and screen the mutant strains.
211
3.
This method may result in the selection of unexpected
secondary
mutations
that
can
interfere
with
the
interpretation of results.
4.
It requires a large number of genetic resources such as
mutant strains and wild-type genes. These resources may
not always be readily available.
212
Protein Engineering
Rational Design
Rational design is the process of creating or engineering proteins
with a specific function or property by designing or altering their
amino acid sequence. This process is based on a thorough
understanding of the structure and function of proteins, as well as
the principles of molecular biology and genetic engineering. Rational
design involves making deliberate changes to a protein's structure
and properties using computational and experimental methods to
improve its stability, activity, selectivity, and specificity. This approach
has many potential applications in biotechnology, medicine, and
other fields where proteins play a critical role.
Principle
Protein engineering is the process of creating new or improved
proteins with specific functional properties through the manipulation
of amino acid sequences. The principle of rational design in protein
engineering
involves
using
computational
and
experimental
methods to design and engineer proteins with specific functions.
This process involves understanding the structure and function of
the target protein and using this knowledge to design new
sequences that can improve or modify its activity, specificity, stability,
and other properties. Rational design in protein engineering is an
efficient and effective approach that can be used to create proteins
with specific properties for a wide range of applications, including
biotechnology, medicine, and industry.
213
Procedure
1.
Identify the protein that needs to be engineered. This could
be a protein that has a specific function or is involved in a
disease process.
2.
Determine the three-dimensional structure of the protein
through various methods such as X-ray crystallography or
NMR spectroscopy. This step is important to identify the
regions of the protein that can be modified without affecting
its overall structure.
3.
Identify the specific site(s) on the protein that needs to be
modified to achieve the desired function. This could be an
active site or a binding site.
4.
Choose the appropriate design strategy based on the
specific requirements. Rational design strategies can be
broadly classified into two types: sequence-based and
structure-based. The former involves modifying the amino
acid sequence of the protein while the latter involves
modifying the protein structure through the introduction of
new amino acids or modification of existing ones.
5.
Design the protein variant: Use appropriate software tools to
design the protein variant. This step involves identifying the
amino acid substitutions or modifications that need to be
made and predicting their effect on the protein structure
and function.
6.
Test the protein variant in vitro using various assays to
determine its function and efficacy. This step could involve
measuring the binding affinity or enzymatic activity of the
variant.
7.
Optimize the protein variant by fine-tuning the amino acid
substitutions or modifications to achieve the desired
214
function. This step could involve iterative rounds of testing
and optimization.
8.
Validate the protein variant in vivo using appropriate animal
models or cell lines. This step is important to determine the
safety and efficacy of the protein variant.
9.
Once the
production
protein variant is
of
the
variant
validated, scale-up the
for
clinical
use
or
commercialization. This step could involve developing a
production process that is cost-effective and scalable.
Advantages
1.
Rational design allows for precise control of the amino acid
sequence and 3D structure of the protein.
2.
It can increase the stability of a protein, allowing it to
withstand harsh environmental conditions or chemical
treatments.
3.
Rational design can enhance the activity of a protein, making
it more effective in its intended function.
4.
It can tailor the properties of a protein to specific
applications, such as drug delivery, biosensors, or industrial
processes.
5.
Rational design can reduce the time and costs associated
with traditional protein engineering methods, such as
directed evolution.
Disadvantages
1.
Rational design relies on a thorough understanding of the
structure and function of the protein, which may be limited
for some proteins.
2.
It can be complex and require advanced knowledge of
protein structure and function.
215
3.
It may not always result in a successful outcome, as
predicting the effects of mutations can be challenging.
4.
It can be time-consuming, especially if multiple rounds of
design and testing are necessary to achieve the desired
properties.
5.
It may limit the diversity of protein variants, as it is focused
on specific changes to the protein sequence.
216
Stem Cell Therapy
DNA Stem Cell Therapy is a revolutionary breakthrough in medical
science that has the potential to cure diseases that were once
considered incurable. Stem cells are cells that can divide and
differentiate into any type of cell in the body. This capability makes
stem cells an essential tool for medical research, especially for the
treatment of various diseases and conditions. One of the latest
advancements in this field is DNA Stem Cell Therapy, which is
gaining recognition for its potential to cure diseases that were once
considered incurable.
DNA Stem Cell Therapy is a process that involves repairing or
altering the DNA in stem cells to treat a specific disease or condition.
Scientists have discovered that by making changes in the DNA of
stem cells, they can alter their function, allowing them to treat a
range of diseases. For instance, researchers have developed
techniques to convert stem cells into neurons to treat neurological
disorders, like Parkinson's disease, or into heart cells to treat heart
disease. This new technology has the potential to revolutionize the
way we treat diseases, and it has the potential to cure some of the
most debilitating conditions that affect humans.
One of the most significant advantages of DNA Stem Cell Therapy is
its ability to cure diseases at the genetic level. In traditional medical
treatments, drugs are used to manage symptoms, but they do not
cure the underlying cause of the disease. With DNA Stem Cell
Therapy, however, the root cause of the disease can be addressed,
offering a cure to many diseases that were once considered
incurable. This technology has the potential to cure diseases like
217
cancer, heart disease, and many others, making it a revolutionary
breakthrough in medical science.
Another advantage of DNA Stem Cell Therapy is its safety and
effectiveness. Stem cells are natural cells in the body, and as such,
they do not cause any adverse effects. Furthermore, since stem cells
are self-renewing, they can provide a long-lasting cure for the
disease. This makes DNA Stem Cell Therapy a highly sought-after
treatment, as it offers a safe and effective cure for many diseases.
With ongoing research and advancements, it is only a matter of time
before DNA Stem Cell Therapy becomes a mainstream treatment
option for many diseases.
Procedure
1.
Collect the stem cells from the patient or from a donor. This
can be done through a biopsy or from a sample of cord
blood. The collected cells are then isolated and cultured in a
laboratory under sterile conditions.
2.
The isolated stem cells are then multiplied or expanded in
the laboratory. This is usually done by culturing the cells in
special media with specific growth factors and nutrients.
3.
The expanded stem cells are then characterized to determine
their type and quality. This is done using various techniques
such as flow cytometry, immunophenotyping, and molecular
analysis.
4.
The characterized stem cells are then prepared for
therapeutic use. This involves selecting the appropriate cells,
modifying them as needed, and processing them to obtain a
suitable dose for transplantation.
5.
The prepared therapeutic cells are then administered to the
patient either intravenously or directly into the site of injury
or disease.
218
6.
The patient is then monitored for any adverse reactions or
changes in their condition. This is done through regular
check-ups and imaging studies.
7.
The success of the stem cell therapy is evaluated over time
by monitoring the patient's response and comparing it to
the baseline. This is done using clinical and imaging
assessments, and measuring the patient's quality of life and
functional status.
219
Reverse Genetics
Reverse genetics is a technique that involves manipulating the DNA
or RNA sequence of a gene or organism in order to understand its
function or phenotype. This technique involves starting with a known
genetic sequence and then modifying it in some way to observe the
resulting effects on the organism. This approach can be used to
study the function of individual genes or to identify the roles of
specific genetic pathways in biological processes. Reverse genetics is
commonly used in research to create knockout or knockdown
models of genes or to introduce mutations into an organism's
genome to study their effects. It is a powerful tool for studying the
genetic basis of disease and for developing new treatments and
therapies.
Principle
The principle of reverse genetics is to work backward from a genetic
sequence to its functional effects, rather than starting with a
phenotype or observable characteristic and trying to identify the
underlying genetic cause. This approach allows researchers to
identify new genes and pathways involved in disease and other
biological processes, and to develop targeted therapies and
treatments based on this knowledge.
Procedure
1.
Identify the gene sequence you want to modify in the
genome and obtain its DNA sequence.
220
2.
Using molecular biology techniques, clone the target gene
sequence into a plasmid or viral vector.
3.
Introduce the recombinant DNA molecule into the host cells
using transfection or transformation methods.
4.
Screen the transfected or transformed cells to identify those
with the desired genetic modification. You can use molecular
markers, fluorescence, or antibiotic resistance genes to
identify positive clones.
5.
Culture the positive clones to obtain a sufficient amount of
cells for further analysis or experiments.
6.
Analyze the genetic modification of the target gene
sequence by sequencing, PCR, or other molecular biology
techniques.
7.
Observe and measure any changes in the phenotype of the
modified cells or organisms. This could include changes in
growth rate, morphology, or biochemical properties.
8.
Verify that the observed phenotype is caused by the genetic
modification of the target gene sequence, rather than other
factors.
9.
Apply the findings to study the biological function of the
modified gene sequence, or to develop new therapies or
treatments for genetic disorders.
Advantages
1.
Reverse genetics allows researchers to identify the function
of a particular gene. By silencing or knocking out a gene,
researchers can determine what its function is in the
organism.
2.
Reverse genetics allows for targeted gene manipulation. It
enables researchers to manipulate specific genes, enabling
them to study the effect of genetic changes.
221
3.
Reverse genetics is a fast and precise way of analyzing gene
function. It allows for the creation of genetically modified
organisms (GMOs) in a short time frame.
Disadvantages
1.
One of the main concerns with reverse genetics is ethical
considerations. The manipulation of an organism’s genes
can be viewed as tampering with nature and can raise ethical
concerns.
2.
Reverse genetics is a technically challenging process that
requires significant expertise and specialized equipment.
This can make it difficult for some researchers to use this
method effectively.
3.
Manipulating a gene can lead to unintended consequences
that are difficult to predict. This can include changes to the
organism’s overall physiology, which can have negative
effects on its health and wellbeing.
222
Transgenic Technology
Transgenic technology, also known as genetic engineering, is a
process of modifying an organism's genetic makeup by adding or
removing genes. It involves the introduction of foreign genes or
DNA sequences into the genome of an organism, thereby altering its
genetic traits. This technology has the potential to enhance the
production of food, medicines, and other valuable products, as well
as to improve the resistance of crops to pests, diseases, and
environmental stresses. However, it also raises ethical and safety
concerns related to the impact of genetically modified organisms on
human health and the environment.
Transgenic Plants
1.
The first step is to isolate the gene of interest, which will be
used for the transformation process. This gene can be
isolated from another plant or synthesized in a laboratory.
2.
The next step is to construct a vector that will carry the
desired gene into the target plant cell. The most commonly
used vectors are plasmids, which are small, circular DNA
molecules.
3.
Once the vector has been constructed, it needs to be
introduced into the target plant cell. This is done through a
process called transformation, which can be accomplished
through several methods, such as Agrobacterium-mediated
transformation, electroporation, or particle bombardment.
4.
After transformation, the plant cells need to be screened for
the presence of the transgene. This is done through a
223
process of selection, in which cells that have incorporated
the transgene are identified and isolated.
5.
Once the transgenic cells have been isolated, they need to
be grown into mature plants. This is done through a process
of regeneration, which involves culturing the transgenic cells
on nutrient-rich media until they form shoots and roots.
6.
Finally, the transgenic plants are characterized to determine
if the desired trait has been successfully introduced. This is
done through various analytical techniques, such as DNA
sequencing, quantitative PCR, and phenotype analysis.
7.
Once
the
transgenic
plants
have
been
successfully
characterized, they can be tested in field trials to determine
their performance under natural conditions.
Transgenic Animals
Transgenic animals are animals that have had their genetic makeup
altered through the insertion, deletion, or replacement of specific
genes using recombinant DNA technology. These animals can be
created using a variety of methods, such as microinjection, where a
gene is directly inserted into an animal's fertilized egg, or somatic
cell nuclear transfer, where the nucleus of an adult animal's cell is
transferred into an egg that has had its nucleus removed.
Transgenic animals have a wide range of applications in various
fields, such as agriculture, medicine, and biotechnology. In
agriculture, transgenic animals can be created that are more resistant
to disease and can produce more milk, meat, or other products. In
medicine, transgenic animals can be used as models for human
disease and to produce therapeutic proteins. In biotechnology,
transgenic animals can be used to produce enzymes and other
industrial products.
224
One of the most common applications is the production of
transgenic mice, which are widely used as a model organism for
human disease. These mice are genetically engineered to carry a
specific human gene that is associated with a disease, such as cancer
or Alzheimer's disease. These mice can then be used to study the
disease and to test potential therapies. Transgenic animals have also
been used to produce human proteins such as clotting factors, which
are used to treat people with genetic disorders such as Hemophilia.
Although transgenic animals have many potential benefits, there are
also worries about their safety and ethics. Some people think that
modifying animals' genes could lead to unintended harm to the
environment and human health. As a result, it's important that we
should also keep a close eye on its future and make sure it's properly
regulated.
1.
The first step in developing a transgenic animal is to identify
the specific gene of interest that will be used to modify the
animal. This gene can be isolated from a variety of sources
including other animals, plants, or bacteria.
2.
Once the gene has been isolated, it must be cloned into a
plasmid, which is a small circular piece of DNA. This process
ensures that the gene is available in large quantities for
further use.
3.
Then a construct is created, which is a vector that contains
the gene of interest along with regulatory elements that
ensure the gene is expressed in the animal. This construct is
introduced into cells, such as embryonic stem cells.
4.
The next step is to introduce the construct into cells that will
be used to create the transgenic animal. This can be done by
a variety of methods, including microinjection or electroporation.
225
5.
The transgenic cells are then used to create transgenic
animals by a process known as blastocyst injection. In this
process, the cells are injected into an early-stage blastocyst,
which is then implanted into a surrogate mother.
6.
Then screen the offspring of the surrogate mother for
successful transgenic animals. This is done by analyzing the
DNA of the animal to confirm that the gene of interest has
been integrated into the animal's genome.
7.
Finally, analyze the phenotype of the transgenic animals to
determine the effects of the transgene on the animal's
development and behavior. This process includes monitoring
the animal's growth, development, and overall health.
8.
The transgenic animals that are produced can be used for
further research or they can be bred to create a line of
animals that contain the transgene. This requires ongoing
maintenance and management of the animals, including
regular monitoring of their health and development.
226
Buffers and Reagents
ACES Buffer
ACES 0.01 M
NaCl 137 mM
KCl 2.7 mM
Na2HPO4 10 mM
KH2PO4 1.8 mM
Ampicillin
(2S,5R,6R)-6-[(R)-(-)-2-Amino-2-phenylacetamido]3,3-dimethyl-7-oxo-4-thia-1azabicyclo[3.2.0]heptane-2-carboxylic acid.
Bovine Serum Albumin (BSA)
A globular protein composed of 583 amino acids.
Ethidium Bromide
C20H14N2Br2
A fluorescent intercalating agent used for the
detection of nucleic acids in gel electrophoresis.
GelRed
It’s a proprietary formulation - a fluorescent nucleic
acid stain used in gel electrophoresis
Glycine Buffer
Glycine 0.01 M
227
NaCl 137 mM
KCl (2.7 mM)
Na2HPO4 (10 mM)
KH2PO4 (1.8 mM)
HEPES Buffer
HEPES 10 mM
NaCl 137 mM
KCl (2.7 mM
Na2HPO4 10 mM
KH2PO4 1.8 mM
IPTG
Isopropyl-beta-D-thiogalactopyranoside
-
a
chemical used to induce the expression of genes in
bacteria that contain a lac promoter.
Kanamycin
(2S,3R,4R,5R,6S)-6-[(2S,3S,4S,5R)-4-amino-5hydroxy-2-(hydroxymethyl)-3-[[(2S,3S,4S,5R,6R)-5amino-2-(hydroxymethyl)-6-[[(2R,3R,4R,5S,6S)-3,4,5trihydroxy-6-(hydroxymethyl)oxan-2-yl]oxy]-4carbamoyloxy-3-carboxy-4-carbamoyloxy-4,5dihydroxyoxan-2-yl]oxy-6-carbamoyloxyoxan-3yl]oxy]tetrahydro-2-furanone - an aminoglycoside
antibiotic used to select for cells containing a
plasmid with a kanamycin resistance gene.
LB Broth
Peptone
yeast extract
NaCl in a buffered solution
228
a rich medium used for growing bacteria in culture.
MES Buffer
MES 0.01 M
NaCl 137 mM
KCl 2.7 mM
Na2HPO4 10 mM
KH2PO4 1.8 mM
PBS-T
Phosphate-buffered saline (PBS) 0.1 M
Tween-20 0.05%
Phosphate Buffered Saline (PBS)
NaCl 137 mM
KCl 2.7 mM
Na2HPO4 10 mM
KH2PO4 1.8 mM
Ripa Buffer
NaCl 150 mM
Tris-base 50 mM
Nonidet P-40 1%
Sodium deoxycholate 0.5%
SDS 0.1%
Protease inhibitor cocktail 1 tablet per 50 ml of
buffer
SDS Buffer
Sodium dodecyl sulfate (SDS) 0.1%
Sodium chloride (NaCl) 50 mM
Tris-HCl 50 mM, pH 8.0
229
Sodium Dodecyl Sulfate (SDS)
C12H25NaO4S - a detergent used to denature
proteins in gel electrophoresis.
TAE Buffer
Tris-base 40 mM
Acetic Acid 1 M
EDTA 0.01 M
Tris-HCl Buffer
Tris-base 0.01 M
HCl 0.1 M
X-Gal
5-bromo-4-chloro-3-indolyl β-D-galactopyranoside
- a substrate for beta-galactosidase that turns blue
when cleaved, used for detecting gene expression.
230
Some Common Lexes
Agarose Gel Electrophoresis
This is a technique used to separate
DNA fragments based on size. An
agarose gel matrix is used to support
the DNA during the electrophoresis
process.
Agarose
A linear polysaccharide composed of
alternating D-galactose and 3, 6anhydro-L-galactopyranose residues.
Alkaline Phosphatase
A hydrolytic enzyme that cleaves
phosphate groups from nucleotides
and other phosphorylated
compounds.
Ammonium Persulfate
A strong oxidizing agent used as a
catalyst in gel electrophoresis; an
initiator in polymerase chain
reactions.
Ampicillin
A penicillin-based antibiotic used to
select bacterial transformants and
prevent bacterial growth in cultures.
Antifoam
A chemical agent used to prevent
foam formation in cell cultures.
Bacterial strain
A pure culture of bacteria used in
biotechnology.
BamHI
A restriction endonuclease that
cleaves DNA at a specific recognition
231
site.
Bovine Serum Albumin (BSA)
A protein used as a blocking agent in
ELISAs and other immunoassays.
Bromophenol Blue
A pH indicator used in gel
electrophoresis to monitor protein
Buffers
migration.
Buffers are solutions that help
maintain a stable pH during various
stages of the genetic engineering
process. Examples include Tris-HCl
and phosphate-buffered saline (PBS).
Calcium Chloride
A salt commonly used to increase the
stability of enzymes during protein
purification.
Carboxy-X-rhodamine (ROX)
A fluorescent dye used as a reference
standard in real-time PCR
experiments.
Cell culture media
A nutrient-rich solution used to grow
cells in a laboratory.
Cellulase
A hydrolytic enzyme that breaks
down cellulose, used in the
production of biofuels.
Chloramphenicol
An antibiotic used to inhibit bacterial
growth in cell cultures and plasmid
preparations.
Coomassie Brilliant Blue
A dye used in protein quantification
assays.
DAPI (4',6-diamidino-2-
A fluorescent dye used to stain DNA
232
phenylindole)
in fluorescence microscopy.
Dextran
A polysaccharide used as a molecular
weight marker in electrophoresis.
Dithiothreitol (DTT)
A reducing agent used to protect
proteins from oxidation and to
prepare protein samples for analysis.
DTT
Dithiothreitol - a reducing agent used
to break disulfide bonds in proteins.
Ethanol
A solvent used to inactivate enzymes
and sterilize solutions.
Ethidium bromide
This is a DNA-staining dye that
intercalates into the DNA molecule,
causing it to fluoresce under UV light.
Ethidium bromide is often used to
visualize DNA fragments after
agarose gel electrophoresis.
Gelatin
A protein derived from collagen, used
as a coating agent for cell cultures.
Glucose
A sugar used as a carbon source in
cell culture media.
Glycerol
A cryoprotectant used to store yeast
and bacteria.
Guanidinium Thiocyanate
A chaotropic salt used to lyse cells
and solubilize proteins in gene
cloning and sequencing experiments.
HEPES (4-(2-hydroxyethyl)-
A buffering agent used in cell culture
1-piperazineethanesulfonic
media.
acid)
233
Hydrogen peroxide
A chemical used as an oxidizing
agent in experiments.
Isopropyl alcohol
An alcohol used for sterilization.
Isopropyl β-D-1-
A chemical used to induce expression
thiogalactopyranoside
of cloned genes in bacteria.
(IPTG)
KCl (Potassium Chloride)
An electrolyte commonly used to
stabilize enzymes during protein
purification.
L-glutamine
An amino acid commonly used as a
supplement in cell culture media.
Ligases
These are enzymes that join together
the ends of DNA molecules. They are
used to seal the ends of the cleaved
DNA after insertion of the foreign
gene. Examples of ligases include T4
DNA ligase and E. coli DNA ligase.
Loading buffer
A solution containing a tracking dye,
a reducing agent, and a stabilizing
agent used to load samples into gels
in electrophoresis.
Luria broth
A nutrient-rich broth used to grow
bacteria in a laboratory.
Lysis buffer
A solution used to break open cells
for DNA extraction.
Lysogeny broth (LB)
A rich growth medium used to
cultivate bacteria.
Lysozyme
An enzyme that cleaves the
234
peptidoglycan layer of bacterial cell
walls, used in bacterial lysis and
purification of bacterial DNA.
Magnesium sulfate
A chemical used to stabilize cell
membranes.
Magnesium Chloride
An electrolyte commonly used in PCR
(MgCl2)
(Polymerase Chain Reaction) and
other enzyme-based reactions.
N, N-Dimethylformamide
An organic solvent used in chemical
(DMF)
synthesis and purification of
biopharmaceuticals.
Neomycin
An antibiotic used to inhibit bacterial
growth in cell cultures and plasmid
preparations.
Nucleic Acid Gel Stains
Fluorescent dyes used to stain DNA
and RNA in gel electrophoresis.
PCR reagents
Polymerase chain reaction (PCR) is a
method used to amplify specific DNA
sequences. Reagents used in PCR
include Taq polymerase, dNTPs
(deoxynucleoside triphosphates), and
primers.
Phenol
A polar solvent used for extraction of
RNA, DNA and other biomolecules
from tissues.
Phosphate-buffered saline
A buffer solution used as a diluent or
(PBS)
washing solution in many
biochemical assays.
235
Plasmids
These are circular pieces of DNA that
are separate from the chromosomal
DNA in a cell. They can be used as
vectors to introduce foreign DNA
into a target organism. Plasmids
often contain origin of replication
and antibiotic resistance genes.
Polyethylene glycol (PEG)
A polymer used to increase the
efficiency of transformation.
Polyvinyl alcohol (PVA)
A polymer used as a gelling agent in
electrophoresis.
Ponceau S
A protein stain used in
electrophoresis to visualize protein
bands and confirm transfer to
nitrocellulose or PVDF membranes.
Protease Inhibitors
Compounds used to inhibit the
activity of proteases in protein
purification and analysis.
Restriction enzymes
These are enzymes that cut DNA at
specific sequences, and are often
used to cleave DNA in preparation
for insertion of a foreign gene.
Examples include EcoRI, HindIII, and
BamHI. Restriction enzymes are
usually composed of proteins.
Sodium chloride (NaCl)
A salt used to adjust the osmotic
pressure of solutions.
Sodium Dodecyl Sulfate
A detergent used to solubilize
(SDS)
proteins in electrophoresis and
236
western blotting.
Sodium Hydroxide
A strong alkaline solution used in the
preparation of cell lysates and
protein samples.
Sodium Pyruvate
A metabolic intermediate used as a
supplement in cell culture media.
Streptomycin
An antibiotic used to select for
bacterial transformants.
Tris-acetate-EDTA (TAE)
A buffering solution commonly used
Buffer
in gel electrophoresis.
Taq polymerase
A protein composed of 5 subunits - a
heat stable DNA polymerase
commonly used in PCR reactions
Transformation reagents
Transformation is the process of
introducing foreign DNA into a target
organism. Reagents used in
transformation include calcium
chloride, heat shock, and
electroporation.
Tris Buffer
A buffer solution used as the
electrophoresis running buffer in
protein SDS-PAGE electrophoresis.
Triton X-100
A non-ionic detergent used in cell
lysis and protein solubilization.
Tryptic Soy Agar
A commonly used solid growth
medium for bacteria.
Tryptone
A complex protein used as a nitrogen
source in growth media.
237
Yeast extract
A source of vitamins and amino acids
used to support the growth of yeast.
Zymolyase
An enzyme used to isolate yeast from
its surrounding cell wall.
238
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Dr HK Garg