Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
METHODOLOGY ARTICLE
Open Access
Electroablation: a method for neurectomy and
localized tissue injury
José Moya-Díaz1†, Oscar A Peña1†, Mario Sánchez1†, Daniela A Ureta1, Nicole G Reynaert1, Consuelo Anguita-Salinas1,2,
Gonzalo Marín3,4 and Miguel L Allende1*
Abstract
Background: Tissue injury has been employed to study diverse biological processes such as regeneration and
inflammation. In addition to physical or surgical based methods for tissue injury, current protocols for localized
tissue damage include laser and two-photon wounding, which allow a high degree of accuracy, but are expensive
and difficult to apply. In contrast, electrical injury is a simple and inexpensive technique, which allows reproducible
and localized cell or tissue damage in a variety of contexts.
Results: We describe a novel technique that combines the advantages of zebrafish for in vivo visualization of cells
with those of electrical injury methods in a simple and versatile protocol which allows the study of regeneration
and inflammation. The source of the electrical pulse is a microelectrode that can be placed with precision adjacent
to specific cells expressing fluorescent proteins. We demonstrate the use of this technique in zebrafish larvae by
damaging different cell types and structures. Neurectomy can be carried out in peripheral nerves or in the spinal
cord allowing the study of degeneration and regeneration of nerve fibers. We also apply this method for the
ablation of single lateral line mechanosensory neuromasts, showing the utility of this approach as a tool for the
study of organ regeneration. In addition, we show that electrical injury induces immune cell recruitment to
damaged tissues, allowing in vivo studies of leukocyte dynamics during inflammation within a confined and
localized injury. Finally, we show that it is possible to apply electroablation as a method of tissue injury and
inflammation induction in adult fish.
Conclusions: Electrical injury using a fine microelectrode can be used for axotomy of neurons, as a general tissue
ablation tool and as a method to induce a powerful inflammatory response. We demonstrate its utility to studies in
both larvae and in adult zebrafish but we expect that this technique can be readily applied to other organisms as
well. We have called this method of electrical based tissue ablation, electroablation.
Keywords: Axotomy, Neurectomy, Tissue ablation, Regeneration, Inflammation, Zebrafish
Background
Induced and localized tissue injury is a powerful tool
widely used to study biological processes, such as regeneration and inflammation. A wide range of tissue injury
approaches have been employed both in regeneration and
inflammation studies. Injury techniques used previously
include surgical methods, such as transection and crush
injury (reviewed in [1]), laser-based wounding [2-5], two
photon microscopy [6], and nanoknives for microscale
* Correspondence: allende@uchile.cl
†
Equal contributors
1
FONDAP Center for Genome Regulation, Facultad de Ciencias, Universidad
de Chile, Casilla 653, Santiago, Chile
Full list of author information is available at the end of the article
axon cutting [7]. Despite the high accuracy of laser and
two photon-based wounding methods, these techniques
have drawbacks such as requiring sophisticated equipment, and being labor-intensive and time-consuming.
Other tissue injury approaches involve injection of toxins
(reviewed in [8]), tissue specific expression of toxins [9],
and tissue specific expression of an enzyme, which converts a nontoxic prodrug into a cytotoxic agent [10,11].
The main disadvantage of genetic approaches and other
high precision techniques is the fact that in contexts like
inflammation or even regeneration, such experimental
models seem unrealistic considering recent evidence
obtained on single axon axotomy [12].
© 2014 Moya-Díaz et al.; licensee BioMed Central Ltd. This is an Open Access article distributed under the terms of the
Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use,
distribution, and reproduction in any medium, provided the original work is properly credited.
Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
In contrast, electrical injury provides a simple and
inexpensive method for tissue injury, which also allows
modulating the extent of damage produced by modifying
current intensity and number of pulses. Thus, in contrast
to surgery based methodologies, the extent of damage
produced by electrical injury and its reproducibility across
different experiments does not rely entirely on the skill of
researcher, but also on the selected parameters for the
electrical pulses.
Electrical damaging of tissues involves thermal, electroporation, and electrochemical interactions [13]. Once
an electrical pulse is applied to a tissue, all its components are affected, however cell membranes appear to
be the most vulnerable structure. Intense electric fields
drive reorganization of lipids, which results in membrane permeabilization, a process called electroporation.
Depending on many physical variables, electroporation
can be transient or permanent [14]. While electroporation
leads to the formation of transient pores in the cell
membrane, Joule heating destabilizes the entire lipid
bilayer [15]. Since cell membranes are composed of a
lipidic bilayer held together by hydration forces, even
small increments in cell membrane temperature result
in complete loss of lipid bilayer integrity [16]. In addition,
strong electrical fields alter the conformation of membrane proteins, as they undergo conformational changes
in order to orientate their dipole moment in the direction
of the field [13]. This effect, called electroconformational
denaturation, is especially strong in voltage gated ion
channels [17]. Since the force of an electric field drops off
quadratically with distance from the focal point, electrical
injury is a suitable approach circumscribing the damage
to a small area of tissue [18-21].
Electrical injury has been widely used to induce damage
in the adult nervous system [22,23]. However, despite the
advantages offered by electrical injury to investigate the
molecular and cellular basis of regeneration and inflammation in vivo, these studies are hampered in most cases
by the opacity of the tissue. Thus, most of our current
knowledge of these events comes from in vitro assays
and analysis of fixed tissues. The optical transparency
of zebrafish during embryonic and larval stages, and
the availability of transgenic lines labeling different cell
types, make it possible to study the behavior of almost
any cell type during regeneration and inflammation
in vivo. In this animal model, it was discovered that tissue
injury induces the generation of a tissue-scale gradient of
hydrogen peroxide that is required for leukocyte recruitment to wounds [24]. Furthermore, recent work in zebrafish shows that hydrogen peroxide generated upon tissue
injury promotes axon growth after skin damage [12].
Furthermore, zebrafish cells and tissues regenerate robustly, a feature that allows the study of regeneration
and inflammation processes in the living animal.
Page 2 of 13
Our laboratory has been studying cell death, regeneration and inflammation in neuromasts of the posterior
lateral line (pLL) of zebrafish larvae. The zebrafish lateral
line is composed of sensory organs called neuromasts,
which are distributed on the body surface. Neuromasts
consist of a core of mechanosensory hair cells surrounded
by support cells. Hair cells of neuromasts are innervated
by neurons that form the pLL nerve which are localized
in a cranial ganglion just posterior to the ear [25]. One
of our goals involves the study of regeneration in a single neuromast and to examine the relationship between
mechanosensory cell regeneration and innervation by
the pLL nerve. Since previous approaches involved the
use of heavy metals [26] or aminoglycosides [27] that
cause damage by toxicity to cells, all lateral line neuromasts are damaged, hampering the study of localized
damage and regeneration in a single isolated organ.
In the present work, we developed an electrical based
method for neurectomy and tissue ablation, which allowed
us to precisely sever nerves and to ablate small areas of
tissue in a simple and reproducible way in both larvae and
adult fish. The protocol can be adapted to study inflammation induced by damage, degeneration of axons and
cells, and their regeneration after damage. We foresee that
it will also be of utility in other model organisms where
it is of interest to damage a small area of tissue in order
to observe the cellular dynamics that accompany the
wounding and/or regenerative process.
Results
A simple methodology for electrical tissue injury
Our aim was to develop a simple protocol for inducing
localized tissue damage for the study of inflammation
and regeneration of different cell types in zebrafish. We
have taken advantage of the numerous transgenic lines
available that label organs and tissues with fluorescent
proteins, and that these can be readily visualized in the
live zebrafish due to the optical transparency of the
embryos and larvae.
In order to induce tissue damage, we modified procedures for delivering precise current pulses developed previously for iontophoresis of neural transport tracers [28].
In our experimental setup, a precision current source
(Figure 1a, black arrow) was configured to deliver electrical pulses of the desired amperage, and they were
applied to zebrafish larvae with a microelecrode held by
a micromanipulator (Figure 1a, black arrowhead). During
the process, zebrafish are mounted (see below) and visualised under a fluorescence microscope to be able to see
the cells or tissues of interest by virtue of their fluorescent
protein expression (Figure 1a, white arrowhead). This
allows placing the microelectrode tip at the desired location. Since the surface of the microelectrode is completely
insulated with exception of the tip, it is possible to apply a
Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
Page 3 of 13
Figure 1 Electroablation setup and dependence of tissue damage on amperage. (a-c) Experimental setup for electroablation in zebrafish
larvae (b) and adults (c). (a) Current pulses are generated by a precision current source (black arrow), which is connected to the sample by the
microelectrode and a ground wire. The microelectrode is held by a micromanipulator (black arrowhead) and zebrafish are visualized under a
fluorescence microscope (white arrowhead). (b) Close-up view of experimental setup for electroablation in larvae. The microelectrode (white arrow)
enters the agarose through one side and a ground wire is connected to the other side (black arrow). Zebrafish larvae are mounted in a drop of agarose
dissolved in E3 (black arrowhead) in the central depression of the acrylic plate (white arrowhead). (c) Adult zebrafish (black arrowhead) are positioned
in an acrylic plate (white arrowhead) with a larger depression and with a small amount of E3. The microelectrode (white arrow) touches the caudal fin
and the ground wire is immersed in the E3 medium (black arrow). (d-f) Merge of acridine orange (green) and bright field images showing progressive
expansion of acridine orange stain as amperage increases. Pulses of different amperages (5 – 25 μA) were applied for 1 second, and acridine orange
stain was performed 2 hpi. (g) Quantification of acridine orange fluorescence shows a direct relationship with applied amperage. Average fluorescence
intensity was measured within a 50 μm radius surrounding the electroablation site. Data was normalized for each larva by the average fluorescence
intensity measured in an adjacent uninjured area. Values are presented as a normalized average ± SEM from 15–21 larvae per condition, from three
independent experiments. Scale bar, 50 μm.
pulse of current at a single point. The diameter of the microelectrode is less than 1 μm at the tip, which, together
with the positioning offered by the micromanipulator,
minimizes the tissue damage derived from placement of
the microelectrode.
Zebrafish larvae are mounted in a drop of low melting
point agarose dissolved in E3 medium (Figure 1b, black
arrowhead) in an acrylic plate (Figure 1b, white arrowhead) that has a central depression. The microelectrode
enters in the agarose through one side (Figure 1b, white
arrow) while the ground wire is immersed in the agarose
on the opposite side (Figure 1b, black arrow) to generate
an electrical field through the larva. Similarly, adult
zebrafish (Figure 1c, black arrowhead) are positioned in
an acrylic plate with a larger depression (Figure 1c, white
arrowhead), without the need of agarose, and a small
amount of E3 is added to allow current conduction.
The microelectrode is positioned over the target tissue
(Figure 1c, white arrow) and the ground wire is immersed
in the E3 medium (Figure 1c, black arrow).
To analyse the effect of pulse amperage on cell death
in electroablated tissue we applied pulses of different
amperages in the trunk of zebrafish larvae and performed
acridine orange staining. As shown in Figure 1d-f, the application of pulses ranging from 5 μA to 25 μA produced
cell damage in skin cells and progressive destruction of
deeper tissues. Quantification of acridine orange staining
reveals that higher amperages induced death in a larger
number of cells compared with that induced by electroablation at lower amperages (Figure 1g). Thus, pulse
Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
amperage can be used to calibrate the extent of damage
inflicted to tissues by electroablation.
Electroablation as a tool for neurectomy in zebrafish
To test electroablation as a suitable methodology for
neurectomy, we used the TgBAC(neurod:EGFP) transgenic
fish that express the Enhanced Green Fluorescent Protein
(EGFP) in the posterior lateral line (pLL) nerves [29]. The
tip of the microelectrode was brought into contact with
an anesthetized larva embedded in low melting point
agarose and then a current pulse of 17 μA was applied
for 1.5 seconds. Electrical injury based neurectomy of
the pLL nerve had no effect on survival of larvae after
seven days (data not shown). Pulses of lower amperage
(~ 15 μA) were unable to completely sever the pLL axons,
while higher amperages (~ 20 μA) induced extensive
tissue damage. The pulse duration was also important, as
2 second-long pulses elicited aberrant trajectories in regenerated nerves (data not shown). For this reason, proper
calibration of the experimental protocol is recommended
in order to determine the lowest amperage and pulse
times necessary to achieve complete neurectomy in all
larvae while preserving the integrity of surrounding tissues.
Neurectomy by electroablation completely severs all
axons of the pLL nerve leaving a gap of 85 ± 5 μm (n = 10;
three independent experiments) at the site of the lesion
(Figure 2a, arrow). To examine the behavior of pLL nerve
axons after neurectomy, we captured time-lapse images of
the trunk of TgBAC(neurod:EGFP) fish five hours after
pLL nerve electroablation (Figure 2a). The detached
(distal) part of the pLL nerve begins to disintegrate from 8
to 11 hours post injury, hpi (Figure 2a, arrowheads), while
the proximal part of the electroablated nerve retracts
towards the ganglion in a process similar to acute axonal
degeneration. Time-lapse imaging of axotomized larvae at
later times (Figure 2b) shows growth of the pLL nerve
through the site of injury and further caudally once the
remaining fragments are completely cleared. Note that
degeneration of the distal part of the pLL nerve is still
ongoing when the regenerating nerve crosses the site of
axotomy. Regeneration of the pLL nerve is completed
by approximately 25 hpi. Interestingly, the dynamics of
regeneration of an neurectomized pLL nerve by using
electroablation are similar to those reported previously
for two-photon axotomy [30].
Electroablation applied to pLL neuromast injury
Our laboratory is interested in understanding the mechanisms involved in neuromast regeneration [26,31]. Previous studies have shown that mechanosensory hair cells
in neuromasts are able to regenerate, while tools used to
damage neuromast hair cells have been based on exposure
to heavy metals [26] or aminoglicosides [27]. These approaches have proven to be useful for regeneration studies
Page 4 of 13
but they have the drawback that they involve widespread
exposure of other larval tissues to the toxicants. Unlike
chemical based approaches, electroablation allows the
induction of a localized damage to a neuromast, making
it feasible to study single neuromast regeneration.
As a proof of principle for localized single neuromast
ablation, we applied two 8 μA pulses for 2 seconds each
in the L3 pLL neuromast of Tg(cxcr4b:mRFP) transgenic
larvae, in which pLL neuromasts and interneuromastic
cells express the red fluorescence protein [32]. As Figure 3a
shows, intact neuromasts have a rosette-like structure
(Figure 3a, arrowheads) and are connected to each other
by interneuromastic cells (Figure 3a, arrows). Figure 3b
shows the trunk of the same larva shown in Figure 3a
at 8 hpi. Complete disappearance of the L3 neuromast
is observed (Figure 3b, arrowhead) while adjacent neuromasts, L2 and L4, as well as most of the surrounding
interneuromastic cells, remain intact. Neuromast electroablation leaves a 55.8 ± 26.3 μm (n = 20; three independent
experiments) gap among interneuromastic cells.
To further analyze the degree of damage to the surrounding tissues at the injury site, we used a contrast
dye (BODIPY-TR) in Tg(−8.0cldnb:lynGFP) transgenic larvae [33] that express EGFP in all cell types of the lateral
line and epithelial cells of the skin (Figure 4). Optical sections of the electroablated area (Figure 4b), compared to
control fish (Figure 4a), show that electroablation generates tissue disruption that can be detected between the
epidermis and underlying cells up to 28 μm deep, including some muscle cells. In spite of the damage inflicted by
electroablation to cells beyond the lateral line cells, we observed regeneration of the ablated neuromasts (Figure 3c).
While, by 24 hpi, none of the electroablated larvae exhibit
regeneration of damaged neuromasts, by 48 hpi these
values increase to 29.2 ± 2.2% (n = 100; three independent
experiments) and by 72 hpi to 58.2 ± 4.4% (n = 100; three
independent experiments). These results demonstrate that
electroablation can be used to inflict localized damage to
sensory organs and to study their regeneration.
Tissue electroablation as a method to induce inflammation
Since it is well described that tissue injury induces inflammation [34,35], we aimed to apply electroablation as
a suitable method to induce leukocyte recruitment to
focally damaged tissues. We used the neurectomy and
tissue electroablation protocols described above but now
using transgenic lines in which immune cells express
fluorescent proteins. Thus, we could now visualize the
behavior of leukocytes during inflammation and resolution
of inflammation in these contexts. Moreover, we aimed
to quantify the resulting inflammation, as electroablation
could be used for genetic or drug screens that use inflammation as a readout. Our results (see below) indicated that
direct quantification of the number of leukocytes recruited
Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
Page 5 of 13
Figure 2 Neurectomy, degeneration and regeneration of the posterior lateral line (pLL) nerve. (a) Degeneration of a neurectomized pLL
nerve after electrical injury. Transgenic TgBAC(neurod:EGFP) larvae with a labeled pLL nerve were neurectomized by applying a 17 μA pulse of
current for 1.5 seconds. At 5 hpi a neurectomized larva was mounted in agarose for time-lapse imaging for 7 hours (anterior to the left). Fragmentation
of the detached nerve fragment proceeds by a breakdown of the axons into many small sections simultaneously throughout the axotomized nerve
(arrowheads). Note the intact contralateral pLL nerve which is visible (above the electroablated nerve, slightly out of focus) in all the images.
(b) Regeneration of an axotomized pLL nerve. Transgenic TgBAC(neurod:EGFP) larvae was treated and imaged as before. In this case, the neurectomized
larva was mounted for imaging for 6 hours starting at 12 hpi to examine regeneration of the pLL nerve. Note the progressive regeneration of
the pLL nerve by elongation of the remaining axon stumps (arrows) as degeneration of the distal part of the axotomized pLL nerve has concluded.
Scale bar, 50 μm; times in hh:mm:ss.
to the site of nerve and neuromast electroablation were not
feasible due to the great number of leukocytes clustering at
the injury site. Thus, the degree of inflammation was
instead measured as the average intensity of fluorescence
of immune cells in a circular area of 50 μm radius around
the injury site.
To examine the inflammatory response induced by
neurectomy of the pLL nerve, we used a Tg(BACneurod:
EGFP; lyz:DsRED2) compound transgenic line, labeling
the pLL nerve in green [29] and myeloid leukocytes
(mainly neutrophils) in red fluorescence [36], respectively.
In these fish, we observed that pLL nerve neurectomy
induces recruitment of leukocytes to the site of electroablation, forming a cell cluster at the electroablation
site within the first 3 hpi (Figure 5a-c). At 6 hpi, the
number of leukocytes recruited to the site of neurectomy
decreases as inflammation resolution proceeds (Figure 5d).
The number of localized leukocytes further decreases
as degeneration of the neurectomized nerve proceeds
(Figure 5e), and is still decreasing by 12 hpi, when the
regenerating nerve has reached the third neuromast of the
pLL (L3) (Figure 5f). To quantify the degree of inflammation elicited by neurectomy, we acquired images and measured mean red fluorescence intensity in a circular area
centered at the electroablation site (Figure 5b, dotted
circle). As is shown in Figure 5g, myeloid leukocytes
accumulate at the site of neurectomy during the first
3 hours and then their number decreases for the next
9 hours as inflammation resolution takes place. Concomitant with the infiltration of neutrophils into the site of
neurectomy, macrophages are also recruited to the site of
injury during the first 2 hours after damage (Additional
file 1). Compound transgenic fish, Tg(mpeg1:EGFP; neurod:
TagRFP), which harbor macrophages that express GFP
[37] and a pLL nerve labeled with red fluorescent protein,
were subjected to pLL neurectomy by application of a
17 μA pulse for 1.5 seconds. A time series of macrophage infiltration dynamics into the area surrounding
Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
Figure 3 Ablation and regeneration of a single posterior lateral
line neuromast. (a-c) Tg(cxcr4b:mRFP) fish show labeling of posterior
lateral line neuromasts and interneuromastic cells. (a) Lateral view
of 72 hpf control larva showing intact lateral line neuromasts
(arrowheads) connected by interneuromastic cells (arrows). (b) Two
8 μA pulses of 2 seconds each were applied to the L3 posterior lateral
line neuromast. Complete dissapearance of the neuromast’s cells is
observed (arrowhead) while adjacent neuromasts (L2 and L4) and
interneuromastic cells remain intact. (c) The same larva is shown at 48
hpi, showing the regenerated L3 neuromast. Scale bar, 100 μm.
the electroablation site shows accumulation of a large
number of macrophages as a result of the induced damage.
Similarly, it is also possible to examine the inflammatory response induced by the electroablation of a pLL
neuromast. We used Tg(cxcr4b:mRFP; mpx:GFP) compound transgenic fish labeling pLL neuromasts in red
[32] and neutrophils in green fluorescence [38] to study
neutrophil inflammation. In these fish, we observed that
pLL neuromast electroablation induces potent recruitment of neutrophils to the site of electroablation (Figure 6).
Recruited neutrophils form a cell cluster at the site of
electroablation by 2 hours after the injury (Figure 6b).
Seven hours after neuromast electroablation, the number
of leukocytes recruited to the site of damage has decreased, suggesting that resolution of neutrophil-mediated
inflammation has taken place (Figure 6c). In vivo time
lapse imaging of an electroablated larva (Additional file 2),
shows how neutrophils migrate interstitially, mostly from
the caudal hematopoietic tissue (CHT), to the damaged
neuromast. There are also neutrophils migrating from the
dorsal ridge to the site of electroablation. Interestingly,
those neutrophils in the CHT that are close to the damaged neuromast, exhibit directional migration to the
wound, while those neutrophils which are farther away
from the ablation site show low or no motility at all.
Accordingly, quantification of neutrophil-mediated inflammation (Figure 6d) shows a rapid increment in the
number of recruited neutrophils during the first 2 hours
after neuromast electroablation, and a decrease to values
similar to those of control larvae over the next 5 hours.
Electroablation applied to other tissues
We further aimed at applying our electrically based tissue injury method to other tissues and also in adult fish.
Our results with pLL nerve neurectomy led us first to
test electroablation in the central nervous system. We
developed a spinal cord injury protocol which allowed
Page 6 of 13
us to completely sever the spinal cord in zebrafish larvae. At 72 hpf, TgBAC(neurod:EGFP) transgenic fish,
which express EGFP in spinal cord neurons and axons,
were anesthetized and mounted in agarose dissolved in
E3, and a 25 μA pulse of 1 second was applied in their
spinal cord. An intact larva before electroablation is
shown in Figure 7a. Figure 7b shows the same larva
after spinal cord electroablation. This protocol eliminates
nervous tissue leaving a gap (Figure 7b, arrowhead) of
84.92 ± 0.037 μm (n = 40; four independent experiments)
and rendering larvae paralyzed (not shown). The same
larva at 5 dpi is shown in Figure 7c; it exhibits robust
regeneration as new fibers have crossed the ablation site
and the larvae show recovery of mobility (not shown).
Next, we used electroablation to induce superficial tissue injury in adult fish. The application of electroablation
in adults involved the use of acrylic plates larger than
those used with larvae (see Methods). Adult transgenic
fish were anesthetized and positioned in acrylic plates, and
a 25 μA pulse of 2 seconds was applied on their caudal
fin. As done before in larvae, we wished to determine if
localized tissue damage inflicted by electroablation in
caudal fins of adult fish elicited neutrophil recruitment
to the site of injury. Adult Tg(mpx:GFP) transgenic fish,
in which neutrophils are labeled in green, were subjected to electroablation in the caudal fin (Figure 7d-f ).
Before tissue injury, neutrophils are scattered in the
caudal fin (Figure 7d). Local tissue injury was applied
(Figure 7e) and 2 hours later, a cluster of neutrophils
has formed at the lesion site (Figure 7f, arrowhead). In
order to quantify neutrophil recruitment, we acquired
images of electroablated caudal fins in the same transgenic fish immediately after electroablation and two hours
later. Quantification of mean fluorescence intensity in a
circular area with a 100 μm radius surrounding the damage point shows a significant increase (p-value = 0.0361,
paired t test) in electroablated vs control fish (mean
fluorescence, after injury: 3.80 ± 0.51, 2hpi: 15.48 ± 4.60;
n = 9, three independent experiments).
Likewise, we induced tissue damage using two transgenic lines that label other tissues in the caudal fin of
adult fish. Figure 7g shows the caudal fin of a Tg(fli1a:
EGFP) transgenic fish, in which blood vessels are labeled
in green [39]. The electrode was placed precisely on a
vessel to sever it (Figure 7h). The result shows that electroablation allows specific damage to a single blood vessel
(see Figure 7i, inset), leaving a gap of 127.4 ± 13.19 μm
(n = 12; three independent experiments) while adjacent vessels remain intact. Similarly, electroablation allowed neuromast ablation in the caudal fin of adult fish (Figure 7j-l).
The caudal fins of adult sqet20 transgenic fish harbor
green-labeled neuromasts [40], which are distributed
regularly in several lines along the fin (Figure 7j). In
this case, electroablation allowed specific damaging of
Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
Page 7 of 13
Figure 4 Extent of damage inflicted to tissues by neuromast electroablation. Transgenic Tg(−8.0cldnb:lynEGFP) fish, which express EGFP in
lateral line cells and epithelial cells of the skin, were stained with BODIPY. Stained larvae were then mounted in agarose and subjected to
electroablation of the L3 neuromast. Confocal images of electroablated and uninjured control larvae were acquired 20 minutes after injury with 1.5 μm
of separation between z-axis optical slices. (a) Sequential z-axis confocal images of an uninjured larva are shown. The intact neuromast (green) exhibits
a rosette-like structure and BODIPY-TR (red) allows visualization of underlying muscular tissue. (b) Images acquired as in (a) of a larva with an
electroablated neuromast showing local loss of cells in the skin, disconnection of interneuromastic cells, destruction of neuromast cells and
complete loss of the rosette-like structure. BODIPY-TR staining also shows a gap in muscular tissue to a depth of 22.5 μm from the skin surface;
deeper sections appear unperturbed. Values indicate distance froom skin surface towards the inside of the larva in μm. Scale bar, 50 μm.
two adjacent neuromasts (see Figure 7j inset). The microelectrode was positioned on the neuromasts (Figure 7k)
and a 25 μA pulse of 2 seconds was applied, destroying
both neuromasts (Figure 7l inset) with no observable
alterations in adjacent neuromasts. These experiments
illustrate how transgenic lines and electroablation can
be easily combined to induce local damage in a tissue of
interest, allowing the study of regeneration and inflammation in adult fish.
Discussion
We describe an electrical injury method, electroablation,
which will contribute to regeneration and inflammation
research by providing a simple and versatile method for
neurectomy, tissue ablation and inflammation induction.
The transparency of zebrafish larvae and availability of
fluorescent tags in specific tissues in transgenic fish, combined with the advantages of an electrical injury method
(localized and precise tissue damage and modulation of
Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
Page 8 of 13
Figure 5 Neutrophil recruitment induced by nerve neurectomy. (a-f) A larva obtained from a cross of TgBAC(neurod:EGFP) and Tg(lyz:DsRED2)
transgenic fish labels the posterior lateral line nerve in green and leukocytes in red. (a) Control larva showing an intact lateral line nerve and few
leukocytes near the nerve. One hour after lateral line nerve neurectomy (b), leukocytes migrate and accumulate at the lesion site (b and c). From
6 hpi, the number of recruited leukocytes decreases and resolution of inflammation takes place. (d-f). The number of recruited leukocytes continues to
decrease as the lateral line nerve regenerates. (g) Quantification of average fluorescence intensity within a 50 μm radius surrounding the neurectomy
site (panel b, dotted circle) shows leukocyte recruitment and subsequent inflammation resolution. Data are presented as average ± SEM from 17 larvae
per condition from two independent experiments. Comparisons were performed by using a repeated measurement two-way ANOVA, with
Bonferroni’s post test. ***, p < 0.001; ns, p > 0.05. Scale bar (a-f) 100 μm. hpi, hours post-injury; a. u., arbitrary units.
the extent of the damage), will now facilitate in vivo
analysis of regeneration and inflammation. Moreover,
electroablation can also be applied to adult fish, making
it possible to carry out regeneration and inflammation
studies in this context.
While other methodologies allow single cell ablation
with high precision or even to sever a single axon [6],
our approach allows localized tissue damaged without
cell type specificity. We think this can be an advantage
rather than a limitation, since nerve and tissue injury
generally involve non-specific damage to several cell
types. Therefore, we propose electroablation as a relevant model for inducing tissue injury in a way that
reflects all of the non-specific tissue damage related
events. This aspect is critical for regeneration studies,
since it has been reported that tissue regeneration depends on hydrogen peroxide production [12,41], which
is induced by tissue damage but does not occur in response to single cell ablation [12].
Our laboratory is particularly interested in the regeneration of both the pLL nerve and neuromasts. We
believe that electroablation will be of value for the study
of the molecular mechanisms and the roles of different
cell types involved in regeneration in both models. In
spite of the high resistence of nerves and their surrounding
tissues, we show successful pLL nerve neurectomy by
electroablation, as a cheap and simple alternative to
more sophisticated approaches, such as two-photon
axotomy [6]. Similarly to previous studies performed by
two-photon axotomy on the pLL nerve [12] we observed
Wallerian degeneration of the detached axons after
electroablation and, later, pLL nerve regeneration. In
addition to pLL nerve regeneration studies, this methodology will allow the study of the mechanisms involved in
hair cell re-innervation, as well as the role of inervation
on hair cell regeneration in this organ system.
Using electroablation we show, for the first time, the
complete ablation and regeneration of a single neuromast.
Since electroablation allows single neuromast ablation, we
foresee that this method will make it possible to study the
mechanisms involved in single neuromast regeneration as
well as the role of different cell types, such as interneuromastic cells, Schwann cells, neutrophils and macrophages
in this process.
Furthermore, we show that electroablation can be
adapted to be used in adult fish, allowing the study of
regeneration and inflammation in adult animals. This
advantage could be used to investigate the effects of chronic
Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
Page 9 of 13
conditions, such as exposure to toxic compounds, on
regeneration and inflammation, for example.
Other potential applications of electroablation
Figure 6 Neuromast electroablation induces an inflammatory
response. Compound transgenic Tg(cxcr4b:mRFP; mpx:GFP) fish
harboring neutrophils expressing green fluorescent protein and
neuromasts labeled in red were used to study neutrophil inflammation
induced by neuromast electroablation. Two 2 second 8 μA pulses
were applied to induce damage to the pLL neuromast. (a-c) The trunk
of a larva subjected to neuromast electroablation is shown with
anterior to the left. (a) Immediately after electroablation, most
neutrophils are present in the caudal hematopoietic tissue (CHT).
(b) At 2 hpi, a large number of neutrophils have specifically migrated
to the site of damage. (c) By 7 hpi, the number of neutrophils at
the site of electroablation has diminished, suggesting resolution of
inflammation is taking place. (d) Quantification of mean fluorescence
in a 50 μm radius around the site of electroablation (panel a, dotted
circle) as a measure of neutrophil recruitment. Data are presented as
average ± SEM from 12 larvae per condition and two independent
experiments. Comparisons were performed by using a repeated
measurement two-way ANOVA, with Bonferroni’s post test. ***, p < 0.001;
ns, p > 0.05. Scale bar (a-b) 100 μm. hpi, hours post-injury; a. u.,
arbitrary units.
In addition to peripheral neurectomy, neuromast ablation, spinal cord injury, inflammation induction and tissue damage in adult fin tissue, we foresee additional
applications using this methodology. First, electroablation can be modified for use in regeneration studies as a
general strategy for ablation of different tissues. Tissues
and organs where electroablation should be feasible include the liver, olfactory rosettes, blood vessels, muscles,
eyes, brain, and ganglia. Furthermore, electroablation
could be used as a reproducible method for injury to
almost any superficial tissue with the aid of the appropriate transgenic fish labeling the tissue of interest. Our
method should be of use even in deep tissues with the
aid of simple surgery protocols, making it possible to
inflict damage in internal organs in adult fish.
Second, since electroablation allows localized tissue
damage and modulation of the degree of damage, this
approach can be used for inflammation studies. Different
methods have been used to induce inflammation in
zebrafish, which include techniques involving physical
methods, such as manual tail transection [42] and laser
induced wounds [5], or a genetically induced chronic
inflammatory condition [43]. Our approach takes advantage of small microelectrodes and electrical pulses to
induce a localized tissue injury that induces immune cell
recruitment. Thus, the main advantages of this method
for inflammation studies are its robustness and versatility.
Since our methodology relies on electrical pulses, it allows
different degrees of tissue damage, which can be achieved
by means of controlling current intensity, and the number
and duration of the pulses applied. This advantage could
be used to investigate possible relationships between the
extent of tissue damage and the inflammatory response,
and furthermore, to study the mechanisms of resolution
of inflammation involved in each case.
Furthermore, this approach allows repeated localized
tissue damage, a useful approach for the study of the
dynamics of immune cells. Since this method can be easily
combined with in vivo time lapse imaging of immune cells
(Additional file 2) and with cell tracking tools, leukocyte
navigation could be studied in an inflammatory context
in vivo, knowing exactly where the inflammatory signals
will be produced. Thus, questions regarding signal integration, receptor desensitization and immune cell
fate after successive inflammatory stimuli, aspects that
remain unknown, could be addressed with the aid of
this methodology.
Finally, electroablation is a powerful tool for neuroinflammation studies, as it allows nervous system injury
or neurectomy as well as inflammation induction. The
Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
Page 10 of 13
Figure 7 Electroablation applied to spinal cord injury in larvae and tissue damage in the caudal fin of adult fish. (a-c) 72 hpf TgBAC
(neurod:EGFP) transgenic larvae were subjected to spinal cord injury. (a) The intact spinal cord before electroablation is shown. The dotted lines
show the outline of the larva. (b) A 1 second 25 μA pulse was applied in the spinal cord (arrowhead), leaving a gap and rendering larvae unable
to move their tails. (c) The regenerated spinal cord at 5 dpi. (d-i) Damaging of different tissues in caudal fin of adult fish by application of a
2 second 25 μA pulse of current. (d-f) Induction of neutrophil recruitment in the caudal fin of adult TgBAC(mpx:GFP) transgenic fish. A large
number of neutrophils recruited to the site of damage is observed by 2 hpi (f). (g-i) Tg(flia:EGFP) transgenic adult fish in which blood vessels are
labeled in green were subjected to electroablation in a single blood vessel (h), leaving a gap in it, as shown in (i) (see inset for higher magnification).
(j-l) Caudal fin neuromast electroablation in sqet20 transgenic fish, which possess neuromasts labeled in green. (j) Intact fin neuromasts are shown.
Within the line of five neuromasts shown in the inset, note positioning of the microelectrode over the second pair of neuromasts (k). (l) After
electroablation, the pair of neuromasts has been destroyed and a gap in the GFP pattern can be observed at that position (see inset for details). Note
that the electrode is easily observable during electroablation facilitating its positioning (e, h and k). Scale bars: a-c, 100 μm; d-l, 500 μm; and i, j, and l
insets 100 μm. dpi, days post-injury.
aforementioned advantages of this approach could be
used to investigate the molecular mechanisms of immune
system involvement in axon or neural regeneration.
Conclusions
In conclusion, electroablation is a versatile tool that will facilitate the study of regeneration and inflammation in a variety of tissues in larvae and adult fish, as well as providing a
simple model for neurectomy and new tools to investigate
regeneration, dynamic aspects of leukocyte recruitment to
wounds, and immune system involvement in regeneration.
Methods
Zebrafish husbandry and experimental conditions
Zebrafish (Danio rerio) larvae were obtained in our facility
according to standard procedures [44]. We used the
transgenic strains TgBAC(neurod:EGFP)nl1 [28], Tg
(neurod:TagRFP)w69 [45], Tg(cxcr4b:mRFP)ump1 [32],
Tg(−8.0cldnb:lynGFP)zf106 [33], Tg(lyz:DsRED2)nz50 [36],
Tg(mpx:GFP)i114 [38], Tg(mpeg1:EGFP) [37], Tg(fli1a:
EGFP)y1 [39] and Et(krt4:EGFP)sqet20 herein named
sqet20 [40]. All embryos were collected by natural
spawning and raised at 28.5°C in E3 medium (5 mM NaCl,
0.17 mM KCl, 0.33 mM CaCl2, 0.3 mM MgSO4, and 0.1%
methylene blue, equilibrated to pH 7.0) in Petri dishes.
Pigment formation was avoided by supplementing E3
medium with PTU (Sigma) 3% from 24 hpf. Embryos
and larvae were staged according to Kimmel et al. [46],
and larval ages are expressed in hours post-fertilization
(hpf ). All animals used in this work were anesthetized
with MS-222 (tricaine, A5040, Sigma, St. Louis, MO,
USA) before each experiment. All procedures complied
with guidelines of the Animal Ethics Committee of the
University of Chile.
Electroablation system components
The setup for electroablation is composed by a precision
current stimulator (AM System, Model 2100; or Catalog
Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
#51595, Stoelting) that provides current pulses to a
tungsten microelectrode (Catalog #UEWMGGSEBN1M,
FHC Inc., Bowdoin, ME, USA) which is positioned with
a micromanipulator (Narishige M-152) (see Figure 1a).
Larvae are visualized with a fluorescence microscope
(AS LMD, Leica) equipped with 10x lens objective. For
visualization at higher magnifications, objectives of longer focal distance are needed, in order to allow the entry
and movement of the microelectrode between the objective and the sample. The stimulator allowed adjustment of current, and number and duration of current
pulses. Tungsten microelectrodes were insulated with
epoxylite, and its final 120 μm of length were tapered at
an angle ranging from 10 to 15°, so that microelectrodes
were less than 1 μm of diameter at the tip. Since microelectrodes are extremely narrow, they needed some preparation to be used with our micromanipulator. We sanded
down the epoxylite from the thicker part of the microelectrode and then placed it in a needle (0.80 × 40 mm,
Catalog #305167, BD). This allowed us to firmly adjust
the microelectrode in the pipette holder of our
micromanipulator.
Tissue injury protocols
TgBAC(neurod:EGFP)nl1 and Tg(neurod:TagRFP) transgenic larvae were used for pLL nerve neurectomy and
spinal cord injury, and Tg(cxcr4b:mRFP)ump1 and Tg
(−8.0cldnb:lynGFP) fish were used for neuromast electroablation. Larvae were anesthetized using MS-222,
embedded in 0,75 or 1% low melting point agarose (BM0130, Winkler, Spain) dissolved in E3 on acrylic plates
(10.7 cm long × 7 cm wide) with a central depression
and positioned using forceps before the agarose set (see
Figure 1b). Both control and experimental larvae were
mounted in order to avoid any manipulation-derived
bias. Note that agarose concentration is important to
facilitate electroablation, since lower concentrations of
agarose are not strong enough to keep embedded larvae
immobile, and thus hamper the penetration of tissues by
the microelectrode (as occurs, for instance, in spinal
cord electroablation) while, conversely, higher concentrations of agarose make it difficult to get the microelectrode into the agarose.
For pLL nerve neurectomy, the microelectrode was
positioned in the region between the pLL nerve ganglion
and the first neuromast. Neurectomy was carried out
by applying a current pulse of 17 μA for 1.5 seconds.
Immediately after a successful electroablation, a gap can
be seen at the site of ablation in the pLL nerve together
with GFP+ labeled debris, possibly corresponding to axon
fragments. All larvae showing incomplete axotomy are
discarded in order to avoid misleading conclusions from
spared axons.
Page 11 of 13
Single neuromast ablations were performed in the third
neuromast of the pLL (L3) by bringing the microelectrode
into contact with the neuromast and then applying two
8 μA pulses, each for 2 seconds. After current application,
careful examination of each larva was performed in order
to discard those larvae with partial neuromast ablation.
With the exception of the time-lapse experiments, electroablated larvae were left unmounted and kept in E3.
Temperature is extremely important in order to achieve
optimal percentages of regenerated neuromasts with
this protocol. Therefore, appropriate incubation at 28°C
of larvae before and after electroablation is advised for
best results.
For spinal cord injury, larvae were mounted sideways
in 1% agarose to keep them immobilized while the microelectrode penetrates the tissue. The microelectrode was
positioned laterally in the region depicted in Figure 7a
(arrowhead), between the horizontal myoseptum and the
dorsal ridge, and then pushed into the larval tissue until it
reached the spinal cord. Spinal cord injury was achieved
by applying a 25 μA pulse for 1 second. After electroablation all those larvae with incomplete spinal cord injury are discarded, which can be evaluated both by
checking for mobility impairment (caudal to the site of
damage) and by looking for surviving axons under a
fluorescence microscope.
Electroablation in caudal fins of adult fish involved
the use of an acrylic plate with a larger depression (see
Figure 1c). Adult fish were anesthetized, and positioned
on the acrylic plate with a minimal amount of E3 to allow
efficient conduction (see Figure 1c). The microelectrode
was positioned as can be seen in Figure 7e, h and k, and a
25 μA pulse for 2 seconds was applied. Since the entire
process can be done in less than a minute electroablation
does not represent a threat to the survival of adult fish.
In order to determine optimal conditions to achieve
regeneration or to induce inflammation in other applications of electroablation, it is best to attempt tissue ablation and neurectomy at a low power with just one pulse,
and then incrementally increase the power and number
of pulses until the appropriate settings are found.
Quantification of neutrophil-mediated inflammation and
acridine orange staining
Compound Tg(BACneurod:EGFP; lyz:DsRED2), for pLL
nerve neurectomy, Tg(cxcr4b:mRFP; mpx:GFP), for pLL
neuromast ablation, and Tg(mpx:GFP), for adult fin electroablation, transgenic larvae were subjected to electroablation as described in the previous section. In order to
quantify leukocyte infiltration, images were acquired of
the damaged site with a fluorescent stereoscope (Olympus,
model MVX10) and then average fluorescence intensity
was measured in a circular area (50 μm radius for quantifications in larvae, and 100 μm in adult fish) centered at
Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
the site of electroablation by using ImageJ software, version 4.2 [47]. The results are expressed in arbitrary units.
Similarly, for acridine orange quantification, images were
acquired from the electroablated fish using a fluorescence
stereoscope (Olympus MVX10) and average fluorescence
intensity was measured within a 50 μm radius around the
site of electroablation and in an adjacent control area
using ImageJ. Average fluorescence intensity in the damaged area was normalized by the measurement taken at
the control site for each larva.
Page 12 of 13
to pLL neurectomy by application of a 17 μA pulse for 1.5 seconds.
A temporal series of images (only green channel shown) shows
macrophage infiltration into the site of axotomy starting 20 minutes after
electroablation. Scale bar, 200 μm. Times expressed in hh:mm:ss.
Additional file 2: Inflammatory neutrophils recruited towards an
electroablated neuromast. Compound Tg(cxcr4b:mRFP; mpx:GFP)
transgenic fish which have red-labeled pLL neuromasts and green-labeled
neutrophils were subjected to neuromast electroablation and immediately
mounted for imaging for 32 minutes under a fluorescent stereoscope.
Images were captured every 30 seconds in the green channel. Interstitial
migration of neutrophils from the caudal hematopoietic tissue (CHT) to the
damaged neuromasts and also from the dorsal ridge can be observed. Scale
bar, 100 μm. Times are expressed as hh:mm:ss.
Image processing
For imaging, larvae were anesthetized and mounted in
low melting point agarose dissolved in E3. Photographs
were taken with a Zeiss LSM 510 Meta confocal microscope, or an Olympus IX81 fluorescence microscope.
For time-lapse imaging, we used a Zeiss Axiovert 200 M
microscope equipped with a 20X lens objective and an
Axiocam camera (Figure 2) or an Olympus MVX10
fluorescent stereoscope and recorded with a QImaging
digital camera (Additional file 1 and Additional file 2).
All images were processed with Zeiss Axiovision (Carl
Zeiss Microimaging GmbH, Jena, Germany) and ImageJ
software Version 4.2 [45]. As described in Figure 2a, at 5
hpi axotomized transgenic larvae (TgBAC(neurod:EGFP))
were anesthetized and embedded in 1% low melting
point agarose dissolved in E3. Images were captured
every 60 minutes for 7 hours. As described in Figure 2b,
12 hpi axotomized transgenic (TgBAC(neurod:EGFP)) larvae were anesthetized and embedded in 1% low melting
point agarose dissolved in E3 for imaging for a total of
6 hours. Images were acquired every 60 minutes for
6 hours. As described in Additional file 1, compound
transgenic fish (Tg(mpeg1:EGFP; neurod:TagRFP)) were
neurectomized and immediately mounted in 0.75% low
melting point agarose dissolved in E3. Images were captured every 15 seconds during 2 hours. As described in
Additional file 2, larvae were subjected to neuromast electroablation and immediately embedded in 1% low melting
point agarose dissolved in E3. Images were captured every
30 seconds for a total of 32 minutes.
Statistical analysis
Data are presented as mean values ± SEM. Statistical analysis was performed using GraphPad Prism version 5.00
for Windows software (GraphPad Software, San Diego,
CA, USA). The probability level for statistical significance
was p < 0.05.
Additional files
Additional file 1: Recruitment of macrophages to the site of
neurectomy. Compound transgenic fish, Tg(mpeg1:EGFP; neurod:TagRFP),
labeling macrophages in green and the pLL nerve in red, were subjected
Abbreviations
pLL: Posterior lateral line; CHT: Caudal hematopoietic tissue; hpf: Hours
post-fertilization; dpf: Days post-fertilization; hpi: Hours post-injury;
dpi: Days post-injury; GFP: Green fluorescent protein; EGFP: Rnhanced
green fluorescent protein; RFP: Red fluorescent protein.
Competing interests
The authors have declared that no competing interests exist.
Authors’ contributions
JM developed the methodology, carried out nerve regeneration studies,
participated in inflammation and neuromast regeneration studies, and helped
to draft the manuscript. MS developed the neuromast ablation protocol, carried
out neuromast regeneration and inflammation studies and helped to draft the
manuscript. OP carried out inflammation studies, performed the statistical
analysis and drafted the manuscript. DU and NR helped to carry out
inflammation studies. CA carried out spinal cord injury studies. GM
participated in the design of the study and development of methodology.
MA conceived of the study, participated in its design and helped to draft
the manuscript. All authors read and approved the final manuscript.
Acknowledgements
We thank Joao Botelho for helpful discussions, Catalina Lafourcade and
Víctor Guzmán for expert fish care and Florencio Espinoza for technical help.
Zebrafish strains were kindly provided by Alex Nechiporuk, David Raible,
Christine Dambly-Chaudière, Darren Gilmour, Phillip Crosier, Stephen
Renshaw, Graham Lieschke, Brant Weinstein and Vladimir Korzh. This work
was supported by grants to MA from FONDAP (15090007), FONDECYT
(1110275) and ICGEB (CRP/CHI11-01) and to GM from FONDECYT (1110281).
Author details
1
FONDAP Center for Genome Regulation, Facultad de Ciencias, Universidad
de Chile, Casilla 653, Santiago, Chile. 2Departamento de Ciencias Biológicas,
Facultad de Ciencias Biológicas, Universidad Andrés Bello, Santiago, Chile.
3
Laboratorio de Neurobiología y Biología del Conocer, Facultad de Ciencias,
Universidad de Chile, Santiago, Chile. 4Facultad de Medicina, Universidad
Finis Terrae, Santiago, Chile.
Received: 12 August 2013 Accepted: 21 January 2014
Published: 16 February 2014
References
1. Rosenzweig ES, McDonald JW: Rodent models for treatment of spinal
cord injury: research trends and progress toward useful repair. Curr Opin
Neurol 2004, 17(2):121–131.
2. Jones JE, Corwin JT: Regeneration of sensory cells after laser ablation in
the lateral line system: hair cell lineage and macrophage behavior
revealed by time-lapse video microscopy. J Neurosci 1996, 16(2):649–662.
3. Yanik MF, Cinar H, Cinar HN, Chisholm AD, Jin Y, Ben-Yakar A: Neurosurgery:
functional regeneration after laser axotomy. Nature 2004, 432(7019):822.
4. Stramer B, Wood W, Galko MJ, Redd MJ, Jacinto A, Parkhurst SM, Martin P:
Live imaging of wound inflammation in Drosophila embryos reveals key
roles for small GTPases during in vivo cell migration. J Cell Biol 2005,
168(4):567–573.
Moya-Díaz et al. BMC Developmental Biology 2014, 14:7
http://www.biomedcentral.com/1471-213X/14/7
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
Redd MJ, Kelly G, Dunn G, Way M, Martin P: Imaging macrophage
chemotaxis in vivo: studies of microtubule function in zebrafish wound
inflammation. Cell Motil Cytoskel 2006, 63(7):415–422.
O’Brien GS, Rieger S, Martin SM, Cavanaugh AM, Sagasti A: Two-photon
axotomy and time-lapse confocal imaging in live zebrafish embryos.
J Vis Exp 2009, 16(24):1129. doi:10.3791/1129.
Chang WC, Hawkes E, Keller CG, Sretavan D: Axon repair: surgical application
at a subcellular scale. WIREs Nanomed Nanobiotechnol 2010, 2(2):151–161.
Wiley RG, KlineIV RH: Neuronal lesioning with axonally transported toxins.
J Neurosci Meth 2000, 103:73–82.
Li Z, Korzh V, Gong Z: DTA-mediated targeted ablation revealed
differential interdependence of endocrine cell lineages in early
development of zebrafish pancreas. Differentiation 2009, 78(4):241–252.
Curado S, Anderson RM, Junqblut B, Mumm J, Schroeter E, Stainier DY:
Conditional targeted cell ablation in zebrafish: a new tool for
regeneration studies. Dev Dyn 2007, 236(4):1025–1035.
Pisharath H, Rhee JM, Swanson MA, Leach SD, Parsons MJ: Targeted
ablation of beta cells in the embryonic zebrafish pancreas using
E. coli nitroreductase. Mech Dev 2007, 124(3):218–229.
Rieger S, Sagasti A: Hydrogen peroxide promotes injury-induced
peripheral sensory axon regeneration in the zebrafish skin. PLoS Biol
2011, 9(5):e1000621.
Lee RC, Zhang D, Hanning J: Biophysical injury mechanisms in electrical
shock trauma. Annu Rev Biomed Eng 2000, 02:477–509.
Lee RC, Canaday DJ, Hammer SM: Transient and stable ionic
permeabilization of isolated skeletal muscle cells after electrical shock.
J Burn Care Rehabil 1993, 14(5):528–540.
Lee RC, Dougherty W: Electrical injury: mechanisms, manifestations, and
therapy. IEEE T Dielect El In 2003, 10(5):810–818.
Gershfeld NL, Murayama M: Thermal instability of red blood cell
membrane bilayers: temperature dependence of hemolysis. J Memb Biol
1988, 101(1):62–72.
Chen W, Lee RC: Altered ion channel conductance and ionic selectivity
induced by large imposed membrane potential pulse. Biophys J 1994,
67(2):603–612.
Hussmann J, Zamboni WA, Russell RC, Roth AC, Kucan JO, Suchy H, Bush K,
Bradley T, Brown RE: A model for recording the microcirculatory changes
associated with standardized electrical injury of skeletal muscle. J Surg
Res 1995, 59(6):725–732.
Carmeliet P, Moons L, Stassen JM, De Mol M, Bouché A, van der Oord JJ,
Kockx M, Collen D: Vascular wound healing and neointima formation
induced by perivascular electric injury in mice. Am J Pathol 1997, 150
(2):761–776.
Fan KW, Zhu ZX, Den ZY: An experimental model of an electrical injury to
the peripheral nerve. Burns 2005, 31(6):731–736.
Kusada A, Isogai N, Cooley BC: Electric injury model of murine arterial
thrombosis. Thromb Res 2007, 121(1):103–106.
Chen XY, Wolpaw JR: Ablation of cerebellar nuclei prevents H-reflex
down-conditioning in rats. Learn Mem 2005, 12:248–254.
Gao L, Fei S, Qiao W, Zhang J, Xing H, Du D: Protective effect of chemical
stimulation of cerebellar fastigial nucleus on stress gastric mucosal injury
in rats. Life Sci 2011, 88:871–878.
Niethammer P, Grabher C, Look AT, Mitchison TJ: A tissue-scale gradient of
hydrogen peroxide mediates rapid wound detection in zebrafish.
Nature 2009, 459(18):996–999.
Ghysen A, Dambly-Chaudière C: Development of the zebrafish lateral line.
Curr Opin Neurobiol 2004, 14(1):67–73.
Hernández PP, Moreno V, Olivari FA, Allende ML: Sub-lethal concentrations
of waterborne copper are toxic to lateral line neuromasts in zebrafish
(Danio rerio). Hearing Res 2006, 213(1–2):1–10.
Froehlicher M, Liedtke A, Groh KJ, Neuhauss SC, Segner H, Eggen RI: Zebrafish
(Danio rerio) neuromast: promising biological endpoint linking
developmental and toxicological studies. Aquat Toxicol 2009, 95(4):307–319.
Alheid GF, Carlsen J: Small injections of fluorescent tracers by
iontophoresis or chronic implantation of micropipettes. Brain Res 1982,
235(1):174–178.
Obholzer N, Wolfson S, Trapani JG, Mo W, Nechiporuk A, Busch-Nentwich E,
Seiler C, Sidi S, Söllner C, Duncan RN, Boehland A, Nicolon T: Vesicular
glutamate transporter 3 is required for synaptic transmission in zebrafish
hair cells. J Neurosci 2008, 28(9):2110–2118.
Page 13 of 13
30. Villegas R, Martin SM, O’Donnell K, Carrillo SA, Sagasti A, Allende ML:
Dynamics of degeneration and regeneration in developing zebrafish
peripheral axons reveals a requirement for extrinsic cell types.
Neural Dev 2012, 7:19.
31. Hernández PP, Olivari FA, Sarrazin AF, Sandoval PC, Allende ML:
Regeneration in zebrafish lateral line neuromasts: expression of the
neural progenitor cell marker Sox2 and proliferation-dependent
and -independent mechanisms of hair cell renewal. Dev Neurobiol 2007,
67:637–654.
32. Gamba L, Cubedo N, Lutfalla G, Ghysen A, Dambly-Chaudière C: Lef1
controls patterning and proliferation in the posterior lateral line
system of zebrafish. Dev Dyn 2010, 239(12):3163–3171.
33. Haas P, Gilmour D: Chemokine signaling mediates self-organizing tissue
migration in the zebrafish lateral line. Dev Cell 2006, 10:673–680.
34. White ES, Mantovani AR: Inflammation, wound repair, and fibrosis:
Reassessing the spectrum of tissue injury and resolution. J Pathol 2013,
229(2):141–144.
35. Pearse D, Jarnagin K: Abating progressive tissue injury and preserving
function after CNS trauma: The role of inflammation modulatory
therapies. Curr Opin Investig Drugs 2010, 11(11):1207–1210.
36. Hall C, Flores MV, Storm T, Crosier K, Crosier P: The zebrafish lysozyme
C promoter drives myeloid-specific expression in transgenic fish.
BMC Dev Biol 2007, 7:42.
37. Ellett F, Pase L, Hayman JW, Andrianopoulos A, Lieschke GJ: mpeg1
promoter transgenes direct macrophage-lineage expression in zebrafish.
Blood 2011, 117(4):e49–e56.
38. Renshaw SA, Loynes CA, Trushell DM, Elworthy S, Ingham PW, Whyte MK:
A transgenic zebrafish model of neutrophilic inflammation. Blood 2006,
108(13):3976–3978.
39. Lawson ND, Weinstein BM: In vivo imaging of embryonic vascular
development using transgenic zebrafish. Dev Biol 2002, 248(2):307–318.
40. Parinov S, Kondrichin I, Korzh V, Emelyanov A: Tol2 transposon-mediated
enhancer trap to identify developmentally regulated zebrafish genes
in vivo. Dev Dyn 2004, 231(2):449–459.
41. Yoo SK, Freisinger CM, LeBert DC, Huttenlocher A: Early redox, Src family
kinase, and calcium signaling integrate wound responses and tissue
regeneration in zebrafish. J Cell Biol 2012, 199(2):225–234.
42. Lieschke GJ, Oates AC, Crowhurst MO, Ward AC, Layton JE: Morphologic
and functional characterization of granulocytes and macrophages in
embryonic and adult zebrafish. Blood 2001, 98(10):3087–3096.
43. Mathias JR, Dodd ME, Walters KB, Rhodes J, Kanki JP, Look AT, Huttenlocher
A: Live imaging of chronic inflammation caused by mutation of
zebrafish Hai1. J Cell Sci 2007, 120(Pt 19):3372–3383.
44. Westerfield M: The Zebrafish Book: A Guide for the Laboratory Use of Zebrafish
(Danio rerio). Eugene, OR: University of Oregon Press; 2000.
45. Mcgraw HF, Snelson CD, Prendergast A, Suli A, Raible DW: Postembryonic
neuronal addition in Zebrafish dorsal root ganglia is regulated by Notch
signaling. Neural Dev 2012, 7:23.
46. Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF: Stages of
embryonic development of the zebrafish. Dev Dyn 1995, 203(3):253–310.
47. Schneider CA, Rasband WS, Eliceiri KW: NIH Image to ImageJ: 25 years of
image analysis. Nat Methods 2012, 9:671–675.
doi:10.1186/1471-213X-14-7
Cite this article as: Moya-Díaz et al.: Electroablation: a method for
neurectomy and localized tissue injury. BMC Developmental Biology
2014 14:7.