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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Semin Cell Dev Biol. Author manuscript; available in PMC 2010 July 1.
Published in final edited form as:
PMCID: PMC2706303

Bioelectric mechanisms in regeneration: unique aspects and future perspectives


Regenerative biology has focused largely on chemical factors and transcriptional networks. However, endogenous ion flows serve as key epigenetic regulators of cell behavior. Bioelectric signaling involves feedback loops, long-range communication, polarity, and information transfer over multiple size scales. Understanding the roles of endogenous voltage gradients, ion flows, and electric fields will contribute to the basic understanding of numerous morphogenetic processes and the means by which they can robustly restore pattern after perturbation. By learning to modulate the bioelectrical signals that control cell proliferation, migration, and differentiation, we gain a powerful set of new techniques with which to manipulate growth and patterning in biomedical contexts. This chapter reviews the unique properties of bioelectric signaling, surveys molecular strategies and reagents for its investigation, and discusses the opportunities made available for regenerative medicine.

Keywords: regeneration, bioelectricity, ion channel, transmembrane potential


Bioelectrical signals are mediated by the steady-state electrical properties of cells and tissues. Despite much fascinating data on the role of endogenous bioelectric signals controlling limb and spinal cord regeneration [13], cell and embryonic polarity [46], growth control [7, 8], and migration guidance of numerous cell types [9], the field as a whole is unknown to several generations of modern cell and developmental biologists. However, some well-known processes, such as the fast, electrical polyspermy block [10, 11], are in fact good examples of such signaling.

This chapter discusses the roles of ion-based physiological processes in guiding cell behavior during regeneration, and more broadly, pattern formation. Functional experiments throughout the last decades showed that some bioelectric events were not merely physiological correlates of housekeeping processes, but rather provided specific instructive signals regulating cell behavior during embryonic development and regenerative repair [12, 13]. Roles for endogenous currents and fields were found in numerous systems (Table 1), and in several cases, spatially-instructive signaling was demonstrated [1419]. Here, I discuss bioelectric controls of morphogenesis in the larger context of pattern formation, outlining controls of individual cell behavior and the unique properties of electrical processes that may underlie the orchestration of higher-order patterning. Specifically excluded in this review are action potentials in neurons, and electromagnetic radiations and biophotons generated by cells. The review concludes with a discussion of the molecular mechanisms transducing bioelectrical events into genetic cascades, and the opportunities provided for the field of regenerative medicine by state-of-the-art molecular tools for the study and manipulation of bioelectric cues.

Table 1
physiological data on endogenous bioelectric signal roles in morphogenesis

Bioelectric signals are generated by specific ion channels and pumps within cell membranes. The segregation of charges achieved by ion fluxes through such transporter proteins gives rise to a transmembrane voltage potential (usually on the order of −50 mV, inside negative). Ion channels and pumps are localized to distinct regions of some cell types; in particular, the apical-basal organization of epithelial cells results in a parallel arrangement of battery cells which in turn gives rise to a transepithelial potential [9, 20]. Thus, all cells – not just excitable neurons and muscle –generate and receive steady-state bioelectrical signals. These transmembrane potentials, electric fields through tissue and surrounding fluids, iso-electric and iso-pH cell groups established by gap junctions [21], and fluxes of individual ions, all carry information to the source cell as well as to its neighbors, and in some cases, to distant locations.

Early discoveries of “animal electricity” can be traced to Luigi Galvani in the late 1700’s, and as early as 1903, it was discovered that hydroids have a specific electrical polarity [22]. However, the majority of the literature in this rich field has come from several subsequent major waves. E. J. Lund, through the 1920’s and 1930’s, focused on currents and showed that polarity was predicted by, and in some cases controlled by, the bioelectric polarity of ion flows in vivo [23]. H. S. Burr (1930’s and 1940’s) focused on measuring and correlating voltage gradients with future developmental pattern in a wide range of species and organs [24, 25]; the measurements suggested that the voltage gradients are quantitatively predictive of morphology, and suggested that the measured fields carried patterning information (an example of Slack’s “second anatomy” [26]). Some of the best early functional results were obtained by Marsh and Beams [15, 27, 28] who were able to specifically control anterior-posterior polarity in planarian regeneration by supplying bioelectrical signals to fragments. Enormously influential for the field was the work of Lionel Jaffe and co-workers including Richard Nuccitelli, Ken Robinson, and Richard Borgens [12, 13, 20, 2937], who demonstrated that electrical properties of individual cells, epithelia, neural structures, and entire limbs were instructive for growth, pattern, and anatomical polarity.

The rise of molecular genetics has drawn attention away from a huge literature of not only descriptive, but also solid, well-controlled functional work using physiological techniques. However, in the last decade, state-of-the-art work has begun to identify proteins responsible for the well-characterized bioelectric signals, the genetic networks that shape them, and the mechanisms that allow cells to transduce the information into growth control decisions. Molecular and cell biology are now being applied to this problem in the areas of wound healing, neural guidance, and cell orientation responses to physiological electric fields [3842], as well as the role of specific ion transporter activity in tail regeneration, left-right patterning, control of adult stem cells and regenerative polarity, and the switch between embryonic stem cell and neoplastic phenotypes [4349].

Although many modern workers are unaware of this rich field, the connection between molecular genetic pathways and bioelectric signaling is being forged by the data itself. A variety of relevant channelopathies have now been discovered by unbiased approaches [50, 51], though ion transporters are usually de-prioritized for analysis when they show up on comparative microarray experiments because it is not yet second nature for cell and molecular biologists to think in terms of bioelectrical signaling. It is hoped that by highlighting the techniques and tools now available, and illustrating strategies for integrating bioelectrical signals with mainstream pathways, workers in multiple sub-fields will consider that modulation of ion flows, currents, and voltages may be at the root of their favorite patterning or mis-patterning problem when ion channels and pumps are identified in genetic screens or subtraction analyses. A superb example of such a convergence is the recent elegant study implicating sodium/hydrogen exchange in planar polarity in Drosophila [52], a relationship that was predicted by bioelectric signals during left-right patterning of embryonic epithelia [53].

Because recent reviews address the role of ionic phenomena and specific ion transporter proteins in wound healing [41, 5456], neoplastic growth [51, 57], and cell cycle [5860], this review has a different goal. Here I will consider the unique properties of bioelectrical signals, as well as the novel techniques being used in this field and the major directions that promise significant advances for regenerative biology and biomedicine [61], both of which require the development of techniques for the rational modulation of 3-dimensional structure at multiple scales.

Context: bioelectric signals as a component of the morphogenetic field

One way to view regeneration of complex structures is as an example of morphostasis - the maintenance of “target morphology” by an organism. This is the shape, defined on multiple scales of size and levels of organization, which a biological system acquires during development, and maintains against cellular turnover (aging), stresses of life (remodeling and wound healing), and major injury requiring regeneration. This is a perspective, focused on information processing in cells and tissues, which emphasizes mechanisms common to the patterning events that occur during embryonic development and regeneration, or fail to occur during neoplastic growth.

The target morphology can be analyzed via mathematical tools - formalized descriptors allowing comparisons of form and of shape transformation, as well as analyses of complexity [26, 6265]. Its presence is revealed not only through highly-stereotypical outcome of embryonic self-assembly, but also in the morphological remodeling over time, observed in both vertebrate and invertebrate systems where deviations from normal shape are slowly corrected. Examples of patterning driven by non-local morphogenetic information include allometric scaling during whole-body remodeling in planaria [66] and the long-term transformation of a tail into a limb when a tail blastema is grafted at the flank in amphibia. Although the origin of blastema cells is local to the site of injury [67] and the initial pattern formation is determined by the original position of the blastema within the donor, the host’s morphogenetic fields exert their influence remotely, and slowly transform the ectopic tail into a limb - the structure appropriate to the large-scale global context in which the blastema is placed [6871].

The mediator of pattern formation and remodeling can be viewed as a “morphogenetic field” [7274] – the sum total of local and long-range patterning signals that impinge upon cells and bear instructive information that orchestrates cell behavior into the maintenance and formation of complex 3-dimensional structures (Fig. 1). While this is currently studied with respects to gradients of chemical messengers [7577], bioelectric signals are also ideal mediators of distributed, non-local field properties in large-scale patterning.

Figure 1
morphogenetic fields and biomedicine

The morphogenetic field, while a classical concept [7881], has recently been reinvigorated through the discovery and molecular characterization of several long-range patterning systems that use the same genetic components to carry patterning signals in embryonic development and regeneration [82, 83]; it is this same information that may be ignored by cells during neoplasm [8486]. This view is a different perspective on regeneration because, rather than focusing on individual molecules and on the special features of regeneration in adults (e.g., scarring), the goal is to understand, and learn to rationally modulate, large-scale patterning processes. This is broadly relevant to many other biomedical areas that can be formulated in terms of establishment, maintenance, and deviation of morphology (e.g., aging, birth defects, and cancer). Examples of underlying mechanisms that establish long-range order are planar cell polarity [8689] and neural signaling. The latter in particular is known to be crucial for regenerative ability [9092], and involved in the maintenance and organization of multicellular structures in the organism such as tongue buds [93], which become disorganized when their innervation is perturbed. This chapter describes one fascinating and molecularly-tractable component of the morphogenetic field: endogenous bioelectric signals.

Cellular-level processes: what can bioelectric signals do?

Coherent regenerative response requires integration of proliferation, cell movement, and differentiation into needed cell types to restore large amounts of organized tissue. Large-scale morphogenesis is the ordered orchestration of lower-level cell behavior, and it is helpful to consider briefly the cell functions that are controlled by endogenous bioelectrical signals.

Cell movement and positioning is an important component of regeneration [94]; movement of progenitor cells towards wounds is observed in planaria [95], zebrafish brain [96], and in mammalian stem cell homing [97]. One of the earliest-observed effects of electric fields on cells was change of orientation (parallel or perpendicular to field lines), growth (extension of processes), or migration (towards the anode or cathode) [98, 99]. Modern protocols avoid artifacts due to polarization of substratum molecules and release of electrode products into medium [100]. Despite some controversy [101] over which cell types respond to physiological-strength electric fields (usually on the order of 50 mV/mm, and as high as 500 mV/mm within the neural tube [102, 103]), it is clear that a large variety of embryonic and somatic cells exhibit galvanotaxis in electric fields of the magnitude often found in vivo [104107]. In embryos, it has been suggested that patterns of voltage gradients form coordinates guiding cell movement during complex morphogenetic processes [108]. Electric guidance also occurs in several types of tumor cells [109]; recently, voltage-gated sodium channels have been strongly implicated in this phenomenon [110, 111] suggesting that endogenous bioelectric states may be a factor in metastatic invasion. It is also now known that bioelectric events are important not only for the generation of guidance signals, but for cell-autonomous responses to fields during migration [112] where channels such as KCa3.1 (KCNN4) provide instructive signals for the direction of cell movement [113].

In addition to cell positioning, regeneration requires the presence of numerous distinct cell types. Early links between ion flow and differentiation were observed by Barth and co-workers, who showed that ventral ectoderm explants could be differentiated into a variety of different cell types by careful modulation of extracellular medium ion content [114, 115]. Bioelectric signals apply not only to embryonic cell differentiation, but also to stem cells, which have unique profiles with respect to ion channel expression and physiological state [116123]. Moreover, it has been recently shown by functional experiments that membrane voltage controls human mesenchymal stem cell (MSC) differentiation in vitro [43]. Much remains to be learned about this process, but it is known for example that Kir2.1 (KCNJ2) channel-mediated hyperpolarization controls differentiation in human myoblasts via a calcineurin pathway [124]. Importantly, a degree of de-differentiation can be induced by ionic modulation [125, 126], and even mature neurons can be coaxed to re-enter the cell cycle by long-term depolarization. This raises the possibility that a degree of stem cell-like plasticity could be induced in terminally-differentiated somatic cells by bioelectric signals [7, 126, 127].

Bioelectric signals also appear to control mitosis rate, which is closely linked to differentiation, as plastic cells tend to proliferate more than most terminally-differentiated somatic cells. Indeed, a comparative analysis of membrane voltage properties of various kinds of cells reveals a striking relationship between depolarization and control of differentiation and proliferation [128]. Numerous studies have implicated K+ currents as protagonists of proliferation and cell cycle progression [129, 130], reviewed in [60, 129]. Cell proliferation appears to be controlled mostly by membrane potential [131, 132], although the effect is not always cell-autonomous: depolarized cells can induce distant neural crest derivatives to over-proliferate [49].

A considerable literature now exists on the role of specific ion transporters, including the sodium-hydrogen exchanger and a variety of K+ and Cl channels, in cell cycle progression, although many questions remain about mechanistic details [129, 133136]. In the zebrafish eye, the V-ATPase is required for retinoblast proliferation [137]. Thus, because of its many patterning roles spanning from the elongation of the tadpole tail [48] to that of pollen tubes [138], as well as in neural stem cells in the regenerating fish brain [139], H+ efflux is a widely relevant transporter for efforts to augment regenerative growth.

The converse of growth through mitosis, that is - cell elimination through programmed cell death, is known to be a part of regeneration in a variety of systems utilizing stem cells [140], tissue renewal [141], and transdifferentiation [142]. Apoptosis is regulated by hyperpolarization via a set of K+ and other channels [59, 143145]; for example, inhibition of K+ channels can promote apoptosis [146148] while activation of K+ channels can inhibit it [149, 150]. Surprisingly, programmed cell death has recently been shown to be required for regeneration [151], suggesting that tight control over programmed cell death (by bioelectric means as well as genetic) may need to be an important aspect of regenerative interventions.

Thus, the data point to transmembrane potential as a broadly-conserved aspect of orchestrating the proliferation, reduction, differentiation, and movement of cells. This is of particular relevance for bioengineers and those seeking to transition findings in regenerative biology into therapeutics: bioelectric events are a powerful, largely-untapped set of cellular control knobs. Gaining the ability to modulate cell number, position, and identity provides the opportunity to manage the alteration or generation of any desired shape.

Higher-level integration: the roles of bioelectric signals in morphogenesis

Use of ion-based signals in higher-order patterning necessitates coupling groups of cells with respect to electrical signals. This often occurs through gap junctions [152, 153], which not only augment cells’ ability to sense extracellular electric fields [154], but also are a common mechanism for organizing cells into functional domains, for example when delimiting regions of neurogenic precursors in the spinal cord [155].

The simplest examples of the roles of ionic signals in multi-cellular systems involve healing epithelial layers, where the fields resulting from disruption of the integrity of the polarized layer provide guidance cues for growth of migratory cells that repair the wound; much molecular data is now available about the alveolar epithelium [156] and the cornea in particular, where not only electric fields [41, 55, 157, 158] but also cell-autonomous changes in transmembrane potential [159, 160] are involved. Other tissues where bioelectric cues contribute to repair include the spinal cord [161163], and indeed this modality is now in human clinical trials with paralyzed patients [164].

A more complex example of morphogenetic control by bioelectric cues is revealed by the role of currents during appendage regeneration. Excellent reviews of the early work of bioelectric effects on regeneration (augmentation of innervation, control of polarity, and alteration of differentiation) are given in [12, 165, 166].

Amputated amphibian limbs maintain a current of injury - a direct-current signal that is very different in regenerating and non-regenerating animals. In the latter, the current decreases slowly as the limb heals, while the former exhibits first a positive polarity (similar to the non-regenerative organism), and then a sharp switch to negative polarity, the peak voltage of which occurs at the time of maximum cell proliferation. For example, in salamanders and newt limbs, which have superb regenerative ability, several hours after amputation the density of stump current density reaches 10–100 mA/cm2 and the electric field is on the order of 50 mV/mm [167]. Currents leave the end of the stump, and re-enter the skin around the limb. The relevant currents can be measured for weeks - much longer than the time needed for the damaged cells to either recover or die, refuting the simple model that the fields reflect passive ion leaks from damaged cells. The studies that correlated changes in voltage and currents were followed by functional experiments. Interfering with the required regeneration gradients via electrical isolation, shunting, ion channel blockers, or exogenous reversal of the gradient inhibited regeneration in several systems [6, 36, 168, 169], demonstrating that these biophysical events were necessary factors regulating regeneration.

Another set of crucial experiments demonstrated sufficiency of the electrical signals in inducing or augmenting regeneration [170, 171]. Guided by measurements of field density, voltage gradient, and direction in endogenous regenerating systems, several labs showed that application of exogenous fields (with physiological parameters) can induce limb regeneration in species which normally do not regenerate, including amphibia [172175], aves [176], and possibly even mammals [177, 178], although the rodent data have not been widely reproduced. For example, when 0.1 mA DC current was artificially pulled out of the stumps of amputated adult Xenopus and Rana forelimbs, treated animals (but not controls) formed broad bifurcated structures [174] containing nerve trunks within the cartilage core and mature epidermal papillae. Cathodal current initiated partial regeneration (including extension of severed ulna, and production of muscle, ligament, and isolated partially segmented cartilage). Implantation of sham electrodes (carrying no current) produced no deviations from the normal response.

Recently, molecular details have been uncovered about the guidance of regenerative events in vertebrate appendages. The tail of Xenopus tadpoles contains spinal cord, muscle, vasculature, and epidermal components. A combination of pharmacological, and molecular-genetic analyses using dominant-negative and constitutively-active ion transporters implicated strong H+ pumping from the wound as an instructive factor in regeneration [48], controlling the appearance of proliferative cells and required for the correct pattern of innervation. Thus, tadpoles normally rely on the V-ATPase hydrogen pump to drive regeneration during early stages. More importantly, during later stages when tadpoles cannot regenerate, the entire regenerative cascade can be reproduced by artificially driving H+ efflux by misexpression of the heterologous (yeast) pump PMA-1 [179].

How are changes in membrane voltage transduced to canonical pathways?

Bioelectric signals are found both upstream and downstream of biochemical and genetic elements (Fig. 2). Ion flows are produced by channels and pumps (which are regulated by transcriptional, translational, and gating mechanisms). Conversely, they control the expression of other genes and the function of physiological mechanisms at the cell surface and in the cytoplasm. Biophysical processes can often achieve considerable patterning in the absence of changes in transcription or even translation, due to the rich regulation of ion transporter activity and the redistribution of macromolecules by electric fields. For example, the stimulation of the sodium-hydrogen exchanger in tumor cell lines results from an increased affinity of the internal H+ regulatory site without changes in expression [180]; likewise, the electrophoretic mechanisms underlying early left-right patterning in frog embryos occur during the first few cleavages, when the zygotic genome is not transcribed [47, 181]. Nevertheless, eventually these processes feed into subsequent pathways that alter gene expression. A commonly-occurring theme of this type is the determination of a cell’s bioelectrical polarity by its anatomical (apical-basal) polarity, which in turn is controlled by the electric fields produced by specifically-localized ion transporters [4, 182184].

Figure 2
integration of bioelectric signals with canonical pathways

Specialized sensory cells can distinguish signals as weak as 5 nV/cm [185, 186]; moreover, these mechanisms can exhibit window effects [187], where a stronger applied signal does not necessarily induce the same effects as a more physiological one. The most common mechanism linking membrane voltage change and downstream events is calcium influx (voltage-sensitive Ca++ channels) [188], though in some instances of K+-dependent signaling, Ca++ fluxes were not affected by K+ channel activity, showing that effects on cell behavior can sometimes bypass modulation of intracellular Ca++ [189]. Additional mechanisms that transduce electrical signal into second-messenger cascades [190] include: modulation of the activity of voltage-sensitive small-molecule transporters (e.g., the serotonin transporter, which converts membrane voltage into the influx of specific chemical signals); redistribution of charged receptors along the cell surface; directional electrophoresis of morphogens through cytoplasmic spaces; and activation of Integrin or other signals by conformational changes in membrane proteins [191193]. These elements can be capitalized upon, for the design of bioelectrical intervention in regenerative processes.

Several more exotic possibilities may be fertile areas for future work. First, it is now clear that the nuclear membrane possesses its own complement of ion transporters, the activity of which expands the relevance of bioelectricity past cell surface events [194] and opens the possibility of specific gene regulation by the membrane potential across nearby nuclear envelope regions. Second, direct changes of specific transcriptional element activity by intracellular potassium ion concentration might mediate ion-specific events independent of membrane voltage per se; this mechanism can involve the DNA-binding activity of such important signaling molecules as p53, forkhead, and CREB (cAMP response element-binding protein) [195]. Third, depolarization has recently been shown to lead to subcellular translocation of NRF-2 transcription factor, providing a mechanistic link between membrane voltage and transcriptional targets [196].

An exciting recent discovery involves VSP – a phosphoinositide phosphatase that converts PI(3,4,5)P3 to PI(4,5)P2 in a manner regulated by a voltage sensor domain [197]. Local levels of PI(4,5)P2 control the cytoskeleton and nuclear effectors. The identification of a protein able to transduce membrane voltage into all of the potential downstream pathways controlled by this powerful second messenger system [198] provides a plethora of testable hypotheses of how membrane depolarization functions in a variety of patterning systems involving migration, apoptosis, and proliferation. Crucially, it was shown that wound healing control by endogenous electric fields is mediated by PTEN [41], adding weight to the possibility that PTEN could be a widely conserved and important means of integrating cell-autonomous ion flows into second messenger and transcriptional responses.

Unique features of bioelectrical signaling processes

Bioelectric networks are essentially recursive. For example, changes in membrane voltage gradients affect the function of voltage-sensitive ion channels, which in turn alters membrane potentials further. Likewise, gap junctions shape electrical properties of cell groups and are themselves sensitive to changes in transmembrane potential and pH. This offers very rich opportunities for biological systems to use ion flows to implement both positive and negative feedback mechanisms. The former, such as that created by the hydrogen/potassium exchanger regulation via potassium-sensitive NF-kB [199], can be used to amplify small physiological signals, while the latter, such as that created by depolarization-induced activation of the hyperpolarizing V-ATPase pump [200], can be used to ensure robustness of patterning against perturbations. For example, consistent left-right patterning is driven by bioelectric cues in early embryogenesis [46, 47, 201203], despite the very significant differences in actual content among normal embryos [204] because the physiological networks can buffer against such genetic and environmental variability.

Bioelectrical signals span several orders of magnitude in scale and levels of organization, controlling the distribution of subcellular components [205, 206] and the structures of epithelia, appendages, and entire embryos [166, 169, 207, 208]. While the penetration of endogenous electrical fields into distant tissue is a function of the complex resistivity and thus often hard to quantify in practice, bioelectric events can exert influence far beyond the local microenvironment. For example, in left-right patterning, a pump-driven battery in ventral cells appears to distribute small molecule morphogens across the entire early frog embryo through long-range gap junction paths under an electrophoretic force [181, 209]. Intriguingly, transplanted tumors can induce large-scale changes in voltage potentials detectable at considerable distances from the primary site [210].

Bioelectrical signals derive some of their behavior from the intrinsic properties governing electric fields, and are an epigenetic mechanism because physiological networks can regulate and generate order in the absence of changes in DNA, RNA, or protein expression. They are likely to be an evolutionarily-ancient example of living systems capitalizing upon “order for free” [211], derived from basic physics which ensures that injury automatically provides cells with a vector cue indicating the position of the damage (Fig. 3). An interesting and important consequence of multi-scale control of bioelectrical signals is their ability to act as “master regulators”: to activate coherent downstream morphogenetic cascades. It has already been shown in physiological experiments that localized interference with signals such as reversal of potential across the neural tube, or shunting specific currents at various anatomical sites, had broad and global effects on patterning [34, 169, 208].

Figure 3
bioelectrical signals leverage the laws of physics into information for living systems

Implications for controlling regeneration

One of the key aspects of understanding signaling in morphogenesis is to ask what information is being carried by a given physiological process and what information capacity the signaling system has. For example, since membrane voltage is only a single parameter, it is likely that the true richness of bioelectrical signaling can only be fully appreciated by considering the microdomains of transporter activity distributed across the entire 2D surface of a cell or epithelium: these inhomogeneities comprise a field of potential values that, because of their spatial distribution, can encode enormous amounts of developmental information [212214]. Although it was appreciated as early as 1983 [215] that individual cells can have more than a single transmembrane potential value, it is still largely unclear how adjacent domains maintain different voltage values and avoid equalizing short-circuits across the underlying cytoplasm. It is likely that we still have a very inadequate picture of all of the bioelectrical signals received and generated by cells in vivo.

With the advent of molecular tools, it is becoming easier to capitalize on this property for augmenting regeneration by providing specific signals. For example, in the case of tail regeneration, a single event – the continuous pumping of H+ at the wound –induces the complete, normal regeneration of the tail. Its patterning and size are correct and its growth is appropriately halted when it catches up with the size of the tails of uncut controls. Two other illustrations are shown in Fig. 4. The ability of relatively simple bioelectrical signals to trigger orchestrated morphogenetic subroutines is a very desirable property for regenerative medicine applications: modulation of physical cues can leverage off the patterning capacity of the host’s genetic programs without needing to micromanage the details of the regenerative process.

Figure 4
sample phenotypes arising from molecular-genetic modulation of bioelectrical cues in Xenopus laevis

The implication of bioelectrical parameters in regulation suggests the idea of the physiological state space, proposed as a hypothesis for guiding future research in this field. Analyses have shown that generally, plastic, embryonic, stem, and tumor cells tend to be depolarized, whereas quiescent terminally-differentiated somatic cells are hyperpolarized [128]. The use of membrane voltage to control cellular state is a powerful tool [49, 216] but it is likely to be only a primitive approximation to the true richness of bioelectrical control. A more useful idea is that cells can be localized in a multi-dimensional physiological state space with a number of orthogonal dimensions indicating membrane voltage, intracellular pH, K+ content, nuclear potential, Cl content, surface charge, etc.

One possibility is that cells can be grouped in distinct regions of this state space corresponding to stem cells, tumor cells, somatic cells, and other types of cells that are of interest to regenerative biology (Fig. 5). This hypothesis implies that in order to make rational changes in cell behavior, 1) data needs to be obtained on multiple cell types from different organs and disease conditions, and 2) strategies need to be developed that use pharmacological reagents targeting natively expressed channels/pumps, and misexpression of well-characterized channel/pump constructs, to move cell states into desired regions (e.g., some cell may need to be depolarized by 30 mV and its internal pH acidified in order to induce proliferation). We are currently using quantitative modeling to expand the XYZTG (3D position, time, and gene expression) space [217] to include the systems biology of bioelectrical properties. The end result of the synthesis of experimental and modeling efforts should be the development of targeted channel/pump modulation strategies to achieve desired bioelectrical states of wound tissues for augmentation of regeneration.

Figure 5
bioelectric state space

One last key aspect of bioelectric signals (Fig. 6) is due to the fact that the same physiological state can be achieved by the function of many different sets of transporters; at the same time, regulatory (e.g., gating) events can result in the same ion transporter functioning very differently in different cells. This disconnect between molecular-genetic profile of cells and bioelectric state is very important: it cannot be assumed that cells expressing the same set of channels and pumps are in the same physiological state. Similarly, comparison of cell types based on microarray or differential expression analysis can be misleading with respect to bioelectric properties. Indeed, knockout of individual channel/pump genes can fail to reveal important aspects of ionic controls because many different transporters can compensate, masking phenotypes. This complexity has a benefit however. For example, in the tadpole, a yeast H+ pump (which does not occur in vertebrates) was used to induce regeneration [48]. It appears that biomedical applications could potentially use any convenient channel or pump to achieve the desired change in cell physiology.

Figure 6
missing the physiological forest for the mRNA/protein expression trees

State-of-the-art tools for research in bioelectric signaling

A variety of new reagents and methodologies have been developed for molecular analysis of bioelectric signals in regenerative contexts [218]. Tools for the characterization of bioelectrical events now include highly-sensitive ion-selective extracellular electrode probes [219, 220], fluorescent reporter dyes, which enable the non-invasive real-time monitoring of pH, membrane voltage, and ion flow in any optically-accessible tissue [221224] (although much opportunity remains for the development of specific, bright, ratiometric dyes that localize exclusively to the desired subcellular locale), and nano-scale voltage reporters [225]. Especially exciting will be the use of multiple physiological dyes in FACS experiments to identify subpopulations of stem and other cell sets that differ in key bioelectric properties, as has been observed for HUVEC cells [226]. Importantly, such experiments on dissociated cells will clearly highlight properties that are cell-autonomous vs. those physiological conditions that can only be maintained as a group phenomenon.

To determine whether ion flow is a causal factor in a particular assay, and to inexpensively and rapidly implicate specific ion transporter proteins for further molecular validation, an inverse drug screen can be performed [227]. This is a chemical genetics approach that capitalizes on a tiered (least-specific → more specific) tree-based distribution of blocker compounds that enables an efficient binary-search for likely candidates. This is most often used to probe endogenous bioelectrical mechanisms and has resulted in the identification of channels and pumps as novel components of left-right patterning [228], anterior-posterior polarity [44], stem cell regulation [43, 45, 49].

It is now possible to use molecular-genetic reagents in gain- and loss-of-function approaches to specifically modulate different aspects of ion flux [49], controlling corneal healing [41], inducing tail regeneration [48] at non-regenerative stages, and drastically altering the positioning and proliferation of neural crest cells [49]. The work of neurobiologists and kidney physiologists has resulted in the availability of a large number of expression constructs encoding ion transporters that can be used as molecular tools for rationally altering the electrical activity of cells and tissues. Morpholino knock-down and mutant/constitutively-active channel and pump construct misexpression are much finer-scale tools than the classical technique of applying current with electrodes, and enable both specific loss-of-function for electrical signals as well as rescue experiments, allowing elegant demonstrations of necessity and sufficiency.

Indeed, analysis of the patterning phenotypes induced by such constructs can be used to dissect the mechanism of action, by distinguishing among different aspects of bioelectrical signals. For example, misexpression of electroneutral transporters can differentiate between the importance of voltage changes vs. that of flux of specific ions. Pore mutants can distinguish between ion conductance roles vs. possible functions of channels/pumps as scaffolds or binding partners (non-electrical signaling); for example, in the Na+/H+ exchanger, both ion-dependent and ion-independent functions control cell directionality and Golgi apparatus localization to wound edge [182]. Gating channel mutants and pumps with altered kinetics can, respectively, be used to reveal upstream signals controlling the bioelectric events, and the temporal properties of the signal. Heterologous transporters, combined with blockade of endogenous channels or pumps, can be used in elegant rescue experiments. Together, these tools can now be used to integrate bioelectrical signals with canonical downstream and upstream pathways, identifying transduction mechanisms leading from ion flow to patterning decisions.

Future Prospects: what’s next?

The field faces a number of major questions. One of the biggest issues is lack of sufficient quantitative data. Many measurements of pH, voltage, and ion content are needed on interesting cell types and model systems to flesh out the physiological state space concept, and compile enough data to develop predictive, quantitative physiological models that encompass the feedback loops and synthesize molecular genetic and bioelectric data [209, 229, 230]. Issues of information content remain a rich area for discovery (what specific messages are encoded for cells by specific kinetics of individual ion fluxes, discrete ranges of transmembrane and trans-epithelial voltage, and distinct regions of different potential throughout the membrane of a single cell?). Oscillations in membrane voltage on a scale much slower than action potentials [231237] are likely to carry important information and must be incorporated into pathway models. Voltage gradients across nuclear and organelle membranes [238] are only beginning to be measured, and their importance for cell function is not yet fully understood [194].

Importantly, even without all of the answers to these many fascinating issues, the existing data provide opportunities for modulation of regeneration. For example, it has been shown that Kv1.3 (KCNA3) and Kv3.1 (KCNC1) blockade increases neural progenitor cell proliferation [239]; likewise, induction of H+ flux induces regeneration of a complex appendage [48] and blockade of gap junction-mediated signals results in the formation of a complete, properly-patterned head in a planarian tail blastema [44]. These techniques can already be integrated into efforts to augment regeneration. The recent development of light-gated ion transporters has been particularly exciting; while these have so far been mainly used for neurobiological studies [240242], they offer the potential of high-resolution spatio-temporal control of bioelectrical changes in cells during regeneration and development.

Three specific directions are being pursued in our group to provide additional opportunities for the field. One is the generation of mutant model species (e.g., Xenopus) expressing fluorescent proteins that report pH [243] or voltage [244], which will greatly augment the ability to study bioelectric properties of cells and tissues in a multitude of regenerative and disease states, or under molecular or pharmacological modulation. Another is the generation of mutants ubiquitously expressing light-gated ion transporters [245247], which will allow unprecedented spatio-temporal control over bioelectric states in any tissue/organ of interest. Finally, in collaboration with bioengineers, we are working on the construction of regenerative sleeves (Fig. 7) –bioreactors to be applied to wounds (e.g., stump amputations) in which the physiological state of wound cells can be precisely controlled by pharmacological, optical, electrical, and genetic means to trigger regeneration and control patterning.

Figure 7
a schematic of the Regeneration Sleeve: application to limb regeneration

The widely-conserved, multi-scale, instructive capacity of bioelectric events, coupled with their ability to induce complex downstream patterning cascades, make ion flow an extremely powerful control modality. Recent discoveries have shed light on the genetic response elements that are activated by ionic signals. The development of specific strategies for modulation of physiological state (whether through gene therapy with controllable transporters or by targeting endogenously-expressed channels), in combination with efforts focused on biochemical factors, is sure to open exciting new vistas in regenerative medicine.


M.L. thanks members of the Levin lab, Richard Nuccitelli, Ken Robinson, Richard Borgens, and Lionel Jaffe for numerous useful discussions, as well as the support of Peter smith and the BRC (NIH P41 RR001395). Ai-Sun Tseng and Dany S. Adams are thanked for their very helpful comments on the manuscript. Fig. 7 was drawn by Jay Dubb; phenotypes in Fig. 4 were obtained by Sherry Aw. M.L. is grateful for support by the NIH (HD055850, GM078484), DARPA (W911NF-07-1-0572) and NHTSA (DTNH22-06-G-00001).


Allometric scaling
remodeling of tissue during changes of cell number or type that maintains correct proportions between organ dimensions (an example of control of nonlocal, large-scale structure)
Bioelectrical signals
information transmitted via spatio-temporal properties of membrane voltage, ion flux, or electrical fields. These are produced by ion channels or pumps functioning in an individual cell or in cell sheets (e.g., epithelial cells arranged in parallel to maximize current) and sensed by the cell itself, neighboring cells, or distant cells. Some are instructive – they carry specific morphogenetic cues used to determine position, differentiation, or proliferation/apoptosis decisions by cells
morphogenesis induced by mechanisms other than changes in DNA sequence or transcription. Bioelectric signals are often epigenetic because these physiological processes can accomplish much patterning via post-translational and physical (e.g., electrophoresis) events not relying on transcription or translation. Ultimately, bioelectric events do induce changes in gene expression
ability of cells to utilize field lines and voltage gradients as migratory cues, moving towards the anode or cathode (depending on cell type)
maintenance, throughout life, of large-scale pattern despite death or injury of individual cells or cell groups
Second anatomy
coding (in terms of positional, gene expression, or signaling factor gradients) of the components of any system. Roughly, this is the molecular identity by which the embryo or regenerating field spatially addresses (maps) its different parts
State space
the set of all possible states of a dynamical system. When applied to cell properties, this is a multi-dimensional theoretical construct where each orthogonal dimension reflects a specific parameter such as voltage, pH, potassium content, etc. Current modeling efforts often make use of the X,Y,Z,t,g space where cells occupy a given point in this space corresponding to their 3-dimensional position, gene expression, etc. We propose a physiological state space that instead groups cells by their bioelectrical properties


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1. Borgens RB. The role of natural and applied electric fields in neuronal regeneration and development. Progress in Clinical & Biological Research. 1986;210:239–50. [PubMed]
2. Borgens RB, Blight AR, Murphy DJ. Axonal regeneration in spinal cord injury: a perspective and new technique. Journal of Comparative Neurology. 1986;250:157–67. [PubMed]
3. Borgens RB, Blight AR, McGinnis ME. Functional recovery after spinal cord hemisection in guinea pigs: the effects of applied electric fields. J Comp Neurol. 1990;296:634–53. [PubMed]
4. Bentrup F, Sandan T, Jaffe L. Induction of Polarity in Fucus Eggs by Potassium Ion Gradients. Protoplasma. 1967;64:254.
5. Novák B, Bentrup FW. An electrophysiological study of regeneration in Acetabularia Mediterranea. Planta. 1972;108:227–44. [PubMed]
6. Novak B, Sirnoval C. Inhibition of regeneration of Acetabularia mediterranea enucleated posterior stalk segments by electrical isolation. Plant Science Letters. 1975;5:183–88.
7. Cone CD, Tongier M. Control of somatic cell mitosis by simulated changes in the transmembrane potential level. Oncology. 1971;25:168–82. [PubMed]
8. Cone CD, Tongier M. Contact inhibition of division: involvement of the electrical transmembrane potential. J Cell Physiol. 1973;82:373–86. [PubMed]
9. McCaig CD, Rajnicek AM, Song B, Zhao M. Controlling cell behavior electrically: current views and future potential. Physiol Rev. 2005;85:943–78. [PubMed]
10. Jaffe LA, Schlichter LC. Fertilization-induced ionic conductances in eggs of the frog, Rana pipiens. J Physiol. 1985;358:299–319. [PubMed]
11. Grey RD, Bastiani MJ, Webb DJ, Schertel ER. An electrical block is required to prevent polyspermy in eggs fertilized by natural mating of Xenopus laevis. Dev Biol. 1982;89:475–84. [PubMed]
12. Borgens R, Robinson K, Vanable J, McGinnis M. Electric Fields in Vertebrate Repair. Alan R. Liss; New York: 1989.
13. Jaffe L. In: Developmental Order: its origin and regulation. Subtelny S, editor. Alan R Liss; New York: 1982. pp. 183–215.
14. Rose SM. Bioelectric Control of Regeneration in Tubularia. American Zoologist. 1974;14:797–803.
15. Marsh G, Beams H. Electrical Control of morphogenesis in regenerating Dugesia tigrina. J Cell Comp Physiol. 1957;39:191–211. [PubMed]
16. Rose SM. Correlation between Bioelectrical and Morphogenetic Polarity During Regeneration in Tubularia. Developmental Biology. 1972;28:274. [PubMed]
17. Dimmitt J, Marsh G. Electrical Control of Morphogenesis in Regenerating Dugesia- Tigrina.2. Potential Gradient Vs Current Density as Control Factors. J Cell Comp Physiol. 1952;40:11–23. [PubMed]
18. Marsh G, Beams HW. Electrical Control of Growth Axis in a Regenerating Annelid. Anatomical Record. 1950;108:512–12.
19. Marsh G, Beams HW. Electrical Control of Morphogenesis in Regenerating Dugesia Tigrina.1. Relation of Axial Polarity to Field Strength. J Cell Comp Physiol. 1952;39:191. [PubMed]
20. Robinson KR. Electric Fields in Vertebrate Repair. Alan R. Liss; 1989. pp. 1–25.
21. Fitzharris G, Baltz JM. Granulosa cells regulate intracellular pH of the murine growing oocyte via gap junctions: development of independent homeostasis during oocyte growth. Development. 2006;133:591–9. [PubMed]
22. Mathews AP. Electrical polarity in the hydroids. Am J Physiol. 1903;8:294–99.
23. Lund E. Bioelectric fields and growth. Univ. of Texas Press; Austin: 1947.
24. Burr HS. The meaning of bioelectric potentials. Yale J Biol Med. 1944;16:353. [PMC free article] [PubMed]
25. Burr HS, Northrop F. The electrodynamic theory of life. Quarterly Review of Biology. 1935;10:322–33.
26. Slack JM. In: Developmental Order: its Origin and Regulation. Subtelny, editor. Alan R. Liss; New York: 1982. pp. 423–36.
27. Marsh G, Beams HW. Electrical Control of Axial Polarity in a Regenerating Annelid. Anatomical Record. 1949;105:513–14.
28. Marsh G, Beams HW. Electrical Control of Growth Polarity in Regenerating Dugesia-Tigrina. Federation Proceedings. 1947;6:163–64. [PubMed]
29. Nuccitelli R. Vibrating Probe - High Spatial-Resolution Extracellular Current Measurement. Federation Proceedings. 1980;39:2129–29.
30. Jaffe LF, Nuccitelli R. An ultrasensitive vibrating probe for measuring steady extracellular currents. J Cell Biol. 1974;63:614–28. [PMC free article] [PubMed]
31. Jaffe LF, Nuccitelli R. Electrical Controls of Development. Ann Rev Biophys Bioengineering. 1977;6:445–76. [PubMed]
32. Jaffe L. The role of ionic currents in establishing developmental pattern. Philosophical Transactions of the Royal Society (Series B) 1981;295:553–66. [PubMed]
33. Nuccitelli R. Endogenous Ionic Currents and DC Electric-Fields in Multicellular Animal-Tissues. Bioelectromagnetics. 1992:147–57. [PubMed]
34. Hotary KB, Robinson KR. Endogenous electrical currents and voltage gradients in Xenopus embryos and the consequences of their disruption. Developmental Biology. 1994;166:789–800. [PubMed]
35. Robinson KR, Messerli MA. In: Nerve Growth and Guidance. McCaig CD, editor. Portland Press; London: 1996. pp. 131–50.
36. Borgens RB. What is the role of naturally produced electric current in vertebrate regeneration and healing. Int Rev Cytol. 1982;76:245–98. [PubMed]
37. Borgens RB. The role of ionic current in the regeneration and development of the amphibian limb. Progress in Clinical & Biological Research. 1983;110(Pt A):597–608. [PubMed]
38. Rajnicek AM, Foubister LE, McCaig CD. Growth cone steering by a physiological electric field requires dynamic microtubules, microfilaments and Rac-mediated filopodial asymmetry. J Cell Sci. 2006;119:1736–45. [PubMed]
39. Rajnicek AM, Foubister LE, McCaig CD. Prioritising guidance cues: directional migration induced by substratum contours and electrical gradients is controlled by a rho/cdc42 switch. Dev Biol. 2007;312:448–60. [PubMed]
40. Zhao M, Dick A, Forrester JV, McCaig CD. Electric field-directed cell motility involves up-regulated expression and asymmetric redistribution of the epidermal growth factor receptors and is enhanced by fibronectin and laminin. Mol Biol Cell. 1999;10:1259–76. [PMC free article] [PubMed]
41. Zhao M, Song B, Pu J, Wada T, Reid B, Tai G, et al. Electrical signals control wound healing through phosphatidylinositol-3-OH kinase-gamma and PTEN. Nature. 2006;442:457–60. [PubMed]
42. Zhao M, Forrester JV, McCaig CD. A small, physiological electric field orients cell division. Proceedings of the National Academy of Sciences of the United States of America. 1999;96:4942–6. [PubMed]
43. Sundelacruz S, Levin M, Kaplan DL. Membrane potential controls adipogenic and osteogenic differentiation of mesenchymal stem cells. PLoS ONE. 2008;3:e3737. [PMC free article] [PubMed]
44. Nogi T, Levin M. Characterization of innexin gene expression and functional roles of gap-junctional communication in planarian regeneration. Dev Biol. 2005;287:314–35. [PubMed]
45. Oviedo NJ, Levin M. smedinx-11 is a planarian stem cell gap junction gene required for regeneration and homeostasis. Development. 2007;134:3121–31. [PubMed]
46. Levin M, Thorlin T, Robinson KR, Nogi T, Mercola M. Asymmetries in H+/K+-ATPase and cell membrane potentials comprise a very early step in left-right patterning. Cell. 2002;111:77–89. [PubMed]
47. Adams DS, Robinson KR, Fukumoto T, Yuan S, Albertson RC, Yelick P, et al. Early, H+-V-ATPase-dependent proton flux is necessary for consistent left-right patterning of non-mammalian vertebrates. Development. 2006;133:1657–71. [PMC free article] [PubMed]
48. Adams DS, Masi A, Levin M. H+ pump-dependent changes in membrane voltage are an early mechanism necessary and sufficient to induce Xenopus tail regeneration. Development. 2007;134:1323–35. [PubMed]
49. Morokuma J, Blackiston D, Adams DS, Seebohm G, Trimmer B, Levin M. Modulation of potassium channel function confers a hyperproliferative invasive phenotype on embryonic stem cells. Proc Natl Acad Sci U S A. 2008;105:16608–13. [PubMed]
50. Stuhmer W, Alves F, Hartung F, Zientkowska M, Pardo LA. Potassium channels as tumour markers. FEBS Lett. 2006;580:2850–2. [PubMed]
51. Arcangeli A, Crociani O, Lastraioli E, Masi A, Pillozzi S, Becchetti A. Targeting ion channels in cancer: a novel frontier in antineoplastic therapy. Curr Med Chem. 2009;16:66–93. [PubMed]
52. Simons M, Gault WJ, Gotthardt D, Rohatgi R, Klein TJ, Shao Y, et al. Electrochemical cues regulate assembly of the Frizzled/Dishevelled complex at the plasma membrane during planar epithelial polarization. Nat Cell Biol. 2009;11:286–94. [PMC free article] [PubMed]
53. Aw S, Levin M. Is left-right asymmetry a form of planar cell polarity? Development. 2009;136:355–66. [PubMed]
54. Nuccitelli R. A role for endogenous electric fields in wound healing. Curr Top Dev Biol. 2003;58:1–26. [PubMed]
55. Huttenlocher A, Horwitz AR. Wound healing with electric potential. N Engl J Med. 2007;356:303–4. [PubMed]
56. Reid B, Song B, McCaig CD, Zhao M. Wound healing in rat cornea: the role of electric currents. Faseb J. 2005;19:379–86. [PMC free article] [PubMed]
57. Kunzelmann K. Ion channels and cancer. J Membr Biol. 2005;205:159–73. [PubMed]
58. Lang F, Foller M, Lang K, Lang P, Ritter M, Vereninov A, et al. Cell volume regulatory ion channels in cell proliferation and cell death. Methods Enzymol. 2007;428:209–25. [PubMed]
59. Lang F, Foller M, Lang KS, Lang PA, Ritter M, Gulbins E, et al. Ion channels in cell proliferation and apoptotic cell death. J Membr Biol. 2005;205:147–57. [PubMed]
60. Wonderlin WF, Strobl JS. Potassium channels, proliferation and G1 progression. J Membr Biol. 1996;154:91–107. [PubMed]
61. Levin M. Large-scale biophysics: ion flows and regeneration. Trends Cell Biol. 2007;17:262–71. [PubMed]
62. Bryant PJ, Bryant SV, French V. Biological regeneration and pattern formation. Scientific American. 1977;237:66–76. [PubMed]
63. French V, Bryant PJ, Bryant SV. Pattern regulation in epimorphic fields. Science. 1976;193:969–81. [PubMed]
64. Barkai N, Ben-Zvi D. ‘Big frog, small frog’--maintaining proportions in embryonic development. Febs J. 2009;276:1196–207. [PubMed]
65. Slack JM. A serial threshold theory of regeneration. J Theor Biol. 1980;82:105–40. [PubMed]
66. Oviedo NJ, Newmark PA, Sanchez Alvarado A. Allometric scaling and proportion regulation in the freshwater planarian Schmidtea mediterranea. Dev Dyn. 2003;226:326–33. [PubMed]
67. Butler EG, O’Brien JP. Effects of localized X-irradiation on regeneration of the Urodele limb. Anatomical Record. 1942;84:407–13.
68. Farinella-Ferruzza N. The transformation of a tail into a limb after xenoplastic transformation. Experientia. 1956;15:304–05.
69. Farinella-Ferruzza N. Risultati di trapianti di bottone codale di urodeli su anuri e vice versa. Riv Biol. 1953;45:523–27. [PubMed]
70. Guyenot E. Le probleme morphogenetique dans la regeneration des urodeles: determination et potentialites des regenerats. Rev Suisse Zool. 1927;34:127–55.
71. Guyenot E, Schotte OE. Greffe de regenerat et differenciation induite. Comptes Rendus de Societe de Phys. His Nat Geneve. 1927;44:21–3.
72. Beloussov LV, Opitz JM, Gilbert SF. Life of Alexander G. Gurwitsch and his relevant contribution to the theory of morphogenetic fields. Int J Dev Biol. 1997;41:771–7. comment 78–9. [PubMed]
73. Martinez-Frias ML, Frias JL, Opitz JM. Errors of morphogenesis and developmental field theory. Am J Med Genet. 1998;76:291–6. [PubMed]
74. Beloussov LV. Morphogenetic fields: outlining the alternatives and enlarging the context. Riv Biol. 2001;94:219–35. [PubMed]
75. Schiffmann Y. The second messenger system as the morphogenetic field. Biochem Biophys Res Commun. 1989;165:1267–71. [PubMed]
76. Schiffmann Y. An hypothesis: phosphorylation fields as the source of positional information and cell differentiation--(cAMP, ATP) as the universal morphogenetic Turing couple. Prog Biophys Mol Biol. 1991;56:79–105. [PubMed]
77. Lewis J. From signals to patterns: space, time, and mathematics in developmental biology. Science. 2008;322:399–403. [PubMed]
78. Driesch H. The science & philosophy of the organism. 2. xii. A. & C. Black; London: 1929. 344 pp.
79. Child CM. Patterns and problems of development. ix. The University of Chicago press; Chicago, Ill: 1941. p. 811.
80. Spemann H. Mrs. Hepsa Ely Silliman Memorial Lectures. [1933] xii. Yale University Press; H. Milford Oxford University Press; New Haven, 1938: Embryonic development and induction; p. 401.
81. Weiss PA. Principles of development; a text in experimental embryology. H. Holt and company; New York: 1939. p. 601.
82. Tataria M, Perryman SV, Sylvester KG. Stem cells: tissue regeneration and cancer. Semin Pediatr Surg. 2006;15:284–92. [PubMed]
83. White RM, Zon LI. Melanocytes in Development, Regeneration, and Cancer. Cell stem cell. 2008;3:242–52. [PubMed]
84. Tsonis PA. Embryogenesis and carcinogenesis: order and disorder. Anticancer Res. 1987;7:617–23. [PubMed]
85. Donaldson DJ, Mason JM. Cancer-related aspects of regeneration research: a review. Growth. 1975;39:475–96. [PubMed]
86. Lee M, Vasioukhin V. Cell polarity and cancer--cell and tissue polarity as a non-canonical tumor suppressor. J Cell Sci. 2008;121:1141–50. [PubMed]
87. Klezovitch O, Fernandez TE, Tapscott SJ, Vasioukhin V. Loss of cell polarity causes severe brain dysplasia in Lgl1 knockout mice. Genes Dev. 2004;18:559–71. [PubMed]
88. Zhan L, Rosenberg A, Bergami KC, Yu M, Xuan Z, Jaffe AB, et al. Deregulation of scribble promotes mammary tumorigenesis and reveals a role for cell polarity in carcinoma. Cell. 2008;135:865–78. [PMC free article] [PubMed]
89. Zallen JA. Planar polarity and tissue morphogenesis. Cell. 2007;129:1051–63. [PubMed]
90. Bryant SV, Fyfe D, Singer M. The effects of denervation on the ultrastructure of young limb regenerates in the newt, Triturus. Developmental Biology. 1971;24:577–95. [PubMed]
91. Maden M. Neurotrophic control of the cell cycle during amphibian limb regeneration. J Embryol Exp Morphol. 1978;48:169–75. [PubMed]
92. Yntema CL. Blastema formation in sparsely innervated and aneurogenic forelimbs of amblystoma larvae. J Exp Zool. 1959;142:423–39. [PubMed]
93. Sollars SI, Smith PC, Hill DL. Time course of morphological alterations of fungiform papillae and taste buds following chorda tympani transection in neonatal rats. J Neurobiol. 2002;51:223–36. [PubMed]
94. Han IS, Seo TB, Kim KH, Yoon JH, Yoon SJ, Namgung U. Cdc2-mediated Schwann cell migration during peripheral nerve regeneration. J Cell Sci. 2007;120:246–55. [PubMed]
95. Salo E, Baguna J. Cell movement in intact and regenerating planarians. Quantitation using chromosomal, nuclear and cytoplasmic markers. J Embryol Exp Morphol. 1985;89:57–70. [PubMed]
96. Zupanc GK. Neurogenesis and neuronal regeneration in the adult fish brain. J Comp Physiol A Neuroethol Sens Neural Behav Physiol. 2006;192:649–70. [PubMed]
97. Chute JP. Stem cell homing. Curr Opin Hematol. 2006;13:399–406. [PubMed]
98. Anderson JD. GALVANOTAXIS OF SLIME MOLD. J Gen Physiol. 1951;35:1–16. [PMC free article] [PubMed]
99. Hyman L, Bellamy A. Studies on the correlation between metabolic gradients, electrical gradients, and galvanotaxis I. Biological Bulletin. 1922;XLIII:313–47.
100. Song B, Gu Y, Pu J, Reid B, Zhao Z, Zhao M. Application of direct current electric fields to cells and tissues in vitro and modulation of wound electric field in vivo. Nature protocols. 2007;2:1479–89. [PubMed]
101. Robinson KR, Cormie P. Electric field effects on human spinal injury: Is there a basis in the in vitro studies? Developmental neurobiology. 2008;68:274–80. [PubMed]
102. Borgens R, Metcalf M, Shi R. Endogenous ionic currents and voltages in amphibian embryos. Journal of Experimental Zoology. 1994;268:307–22.
103. Hotary KB, Robinson KR. The neural tube of the Xenopus embryo maintains a potential difference across itself. Brain Research Developmental Brain Research. 1991;59:65–73. [PubMed]
104. Stump RF, Robinson KR. Xenopus neural crest cell migration in an applied electrical field. J Cell Biol. 1983;97:1226–33. [PMC free article] [PubMed]
105. Zhao M, McCaig CD, Agius-Fernandez A, Forrester JV, Araki-Sasaki K. Human corneal epithelial cells reorient and migrate cathodally in a small applied electric field. Curr Eye Res. 1997;16:973–84. [PubMed]
106. Pullar CE, Isseroff RR. Cyclic AMP mediates keratinocyte directional migration in an electric field. J Cell Sci. 2005;118:2023–34. [PubMed]
107. Nishimura KY, Isseroff RR, Nuccitelli R. Human keratinocytes migrate to the negative pole in direct current electric fields comparable to those measured in mammalian wounds. J Cell Sci. 1996;109 (Pt 1):199–207. [PubMed]
108. Shi R, Borgens RB. Three-dimensional gradients of voltage during development of the nervous system as invisible coordinates for the establishment of embryonic pattern. Dev Dyn. 1995;202:101–14. [PubMed]
109. Yan X, Han J, Zhang Z, Wang J, Cheng Q, Gao K, et al. Lung cancer A549 cells migrate directionally in DC electric fields with polarized and activated EGFRs. Bioelectromagnetics. 2009;30:29–35. [PubMed]
110. Mycielska ME, Djamgoz MB. Cellular mechanisms of direct-current electric field effects: galvanotaxis and metastatic disease. J Cell Sci. 2004;117:1631–9. [PubMed]
111. Fraser SP, Diss JK, Chioni AM, Mycielska ME, Pan H, Yamaci RF, et al. Voltage-gated sodium channel expression and potentiation of human breast cancer metastasis. Clin Cancer Res. 2005;11:5381–9. [PubMed]
112. Schwab A. Function and spatial distribution of ion channels and transporters in cell migration. Am J Physiol Renal Physiol. 2001;280:F739–47. [PubMed]
113. Schwab A, Gabriel K, Finsterwalder F, Folprecht G, Greger R, Kramer A, et al. Polarized ion transport during migration of transformed Madin-Darby canine kidney cells. Pflugers Arch. 1995;430:802–7. [PubMed]
114. Barth LG, Barth LJ. Ionic regulation of embryonic induction and cell differentiation in Rana pipiens. Dev Biol. 1974;39:1–22. [PubMed]
115. Barth LJ, Barth LG. Effect of the potassium ion on induction of notochord from gastrula ectoderm of Rana pipiens. Biol Bull. 1974;146:313–25. [PubMed]
116. Park KS, Jung KH, Kim SH, Kim KS, Choi MR, Kim Y, et al. Functional expression of ion channels in mesenchymal stem cells derived from umbilical cord vein. Stem Cells. 2007;25:2044–52. [PubMed]
117. Flanagan LA, Lu J, Wang L, Marchenko SA, Jeon NL, Lee AP, et al. Unique dielectric properties distinguish stem cells and their differentiated progeny. Stem Cells. 2008;26:656–65. [PubMed]
118. Balana B, Nicoletti C, Zahanich I, Graf EM, Christ T, Boxberger S, et al. 5-Azacytidine induces changes in electrophysiological properties of human mesenchymal stem cells. Cell Res. 2006;16:949–60. [PubMed]
119. Chafai M, Louiset E, Basille M, Cazillis M, Vaudry D, Rostene W, et al. PACAP and VIP promote initiation of electrophysiological activity in differentiating embryonic stem cells. Ann N Y Acad Sci. 2006;1070:185–9. [PubMed]
120. Ravens U. [Electrophysiological properties of stem cells] Herz. 2006;31:123–6. [PubMed]
121. Wenisch S, Trinkaus K, Hild A, Hose D, Heiss C, Alt V, et al. Immunochemical, ultrastructural and electrophysiological investigations of bone-derived stem cells in the course of neuronal differentiation. Bone. 2006;38:911–21. [PubMed]
122. Biagiotti T, D’Amico M, Marzi I, Di Gennaro P, Arcangeli A, Wanke E, et al. Cell renewing in neuroblastoma: electrophysiological and immunocytochemical characterization of stem cells and derivatives. Stem Cells. 2005 E-pub ahead of print. [PubMed]
123. Wang K, Xue T, Tsang SY, Van Huizen R, Wong CW, Lai KW, et al. Electrophysiological properties of pluripotent human and mouse embryonic stem cells. Stem Cells. 2005;23:1526–34. [PubMed]
124. Konig S, Beguet A, Bader CR, Bernheim L. The calcineurin pathway links hyperpolarization (Kir2.1)-induced Ca2+ signals to human myoblast differentiation and fusion. Development. 2006;133:3107–14. [PubMed]
125. Harrington DB, Becker RO. Electrical stimulation of RNA and protein synthesis in the frog erythrocyte. Exp Cell Res. 1973;76:95–8. [PubMed]
126. Cone CD, Cone CM. Induction of mitosis in mature neurons in central nervous system by sustained depolarization. Science. 1976;192:155–8. [PubMed]
127. Stillwell EF, Cone CM, Cone CD. Stimulation of DNA synthesis in CNS neurones by sustained depolarisation. Nat New Biol. 1973;246:110–1. [PubMed]
128. Binggeli R, Weinstein R. Membrane potentials and sodium channels: hypotheses for growth regulation and cancer formation based on changes in sodium channels and gap junctions. J Theor Biol. 1986;123:377–401. [PubMed]
129. MacFarlane SN, Sontheimer H. Changes in ion channel expression accompany cell cycle progression of spinal cord astrocytes. Glia. 2000;30:39–48. [PubMed]
130. Rouzaire-Dubois B, Gerard V, Dubois JM. Involvement of K+ channels in the quercetin-induced inhibition of neuroblastoma cell growth. Pflugers Arch. 1993;423:202–5. [PubMed]
131. Cone CD. The role of the surface electrical transmembrane potential in normal and malignant mitogenesis. Ann NY Acad Sci. 1974;238:420–35. [PubMed]
132. Arcangeli A, Carla M, Bene M, Becchetti A, Wanke E, Olivotto M. Polar/apolar compounds induce leukemia cell differentiation by modulating cell-surface potential. PNAS. 1993;90:5858–62. [PubMed]
133. Valenzuela SM, Mazzanti M, Tonini R, Qiu MR, Warton K, Musgrove EA, et al. The nuclear chloride ion channel NCC27 is involved in regulation of the cell cycle. J Physiol. 2000;529(Pt 3):541–52. [PubMed]
134. Ouadid-Ahidouch H, Ahidouch A. K+ channel expression in human breast cancer cells: involvement in cell cycle regulation and carcinogenesis. J Membr Biol. 2008;221:1–6. [PubMed]
135. Putney LK, Barber DL. Na-H exchange-dependent increase in intracellular pH times G2/M entry and transition. J Biol Chem. 2003;278:44645–9. [PubMed]
136. Boutillier AL, Kienlen-Campard P, Loeffler JP. Depolarization regulates cyclin D1 degradation and neuronal apoptosis: a hypothesis about the role of the ubiquitin/proteasome signalling pathway. Eur J Neurosci. 1999;11:441–8. [PubMed]
137. Nuckels RJ, Ng A, Darland T, Gross JM. The vacuolar-ATPase complex regulates retinoblast proliferation and survival, photoreceptor morphogenesis, and pigmentation in the zebrafish eye. Invest Ophthalmol Vis Sci. 2009;50:893–905. [PubMed]
138. Certal AC, Almeida RB, Carvalho LM, Wong E, Moreno N, Michard E, et al. Exclusion of a proton ATPase from the apical membrane is associated with cell polarity and tip growth in Nicotiana tabacum pollen tubes. Plant Cell. 2008;20:614–34. [PubMed]
139. Zupanc MM, Wellbrock UM, Zupanc GK. Proteome analysis identifies novel protein candidates involved in regeneration of the cerebellum of teleost fish. Proteomics. 2006;6:677–96. [PubMed]
140. Hwang JS, Kobayashi C, Agata K, Ikeo K, Gojobori T. Detection of apoptosis during planarian regeneration by the expression of apoptosis-related genes and TUNEL assay. Gene. 2004;333:15–25. [PubMed]
141. Nadal-Ginard B, Kajstura J, Anversa P, Leri A. A matter of life and death: cardiac myocyte apoptosis and regeneration. J Clin Invest. 2003;111:1457–9. [PMC free article] [PubMed]
142. Mescher AL, White GW, Brokaw JJ. Apoptosis in regenerating and denervated, nonregenerating urodele forelimbs. Wound Repair Regen. 2000;8:110–6. [PubMed]
143. Wang L, Zhou P, Craig RW, Lu L. Protection from cell death by mcl-1 is mediated by membrane hyperpolarization induced by K(+) channel activation. J Membr Biol. 1999;172:113–20. [PubMed]
144. Gilbert MS, Saad AH, Rupnow BA, Knox SJ. Association of BCL-2 with membrane hyperpolarization and radioresistance. J Cell Physiol. 1996;168:114–22. [PubMed]
145. Wang H, Zhang Y, Cao L, Han H, Wang J, Yang B, et al. HERG K+ channel, a regulator of tumor cell apoptosis and proliferation. Cancer Res. 2002;62:4843–8. [PubMed]
146. Chin LS, Park CC, Zitnay KM, Sinha M, DiPatri AJ, Jr, Perillan P, et al. 4-Aminopyridine causes apoptosis and blocks an outward rectifier K+ channel in malignant astrocytoma cell lines. J Neurosci Res. 1997;48:122–7. [PubMed]
147. Miki T, Iwanaga T, Nagashima K, Ihara Y, Seino S. Roles of ATP-sensitive K+ channels in cell survival and differentiation in the endocrine pancreas. Diabetes. 2001;50 (Suppl 1):S48–51. [PubMed]
148. Miki T, Tashiro F, Iwanaga T, Nagashima K, Yoshitomi H, Aihara H, et al. Abnormalities of pancreatic islets by targeted expression of a dominant-negative KATP channel. Proceedings of the National Academy of Sciences of the United States of America. 1997;94:11969–73. [PubMed]
149. Lauritzen I, De Weille JR, Lazdunski M. The potassium channel opener (−)-cromakalim prevents glutamate-induced cell death in hippocampal neurons. J Neurochem. 1997;69:1570–9. [PubMed]
150. Lauritzen I, Zanzouri M, Honore E, Duprat F, Ehrengruber MU, Lazdunski M, et al. K+-dependent cerebellar granule neuron apoptosis. Role of task leak K+ channels. J Biol Chem. 2003;278:32068–76. [PubMed]
151. Tseng AS, Adams DS, Qiu D, Koustubhan P, Levin M. Apoptosis is required during early stages of tail regeneration in Xenopus laevis. Dev Biol. 2007;301:62–9. [PMC free article] [PubMed]
152. Levin M. Gap junctional communication in morphogenesis. Prog Biophys Mol Biol. 2007;94:186–206. [PMC free article] [PubMed]
153. Nicholson BJ. Gap junctions - from cell to molecule. J Cell Sci. 2003;116:4479–81. [PubMed]
154. Cooper MS. Gap junctions increase the sensitivity of tissue cells to exogenous electric fields. J Theor Biol. 1984;111:123–30. [PubMed]
155. Russo RE, Reali C, Radmilovich M, Fernandez A, Trujillo-Cenoz O. Connexin 43 delimits functional domains of neurogenic precursors in the spinal cord. J Neurosci. 2008;28:3298–309. [PubMed]
156. Trinh NT, Prive A, Kheir L, Bourret JC, Hijazi T, Amraei MG, et al. Involvement of KATP and KvLQT1 K+ channels in EGF-stimulated alveolar epithelial cell repair processes. Am J Physiol Lung Cell Mol Physiol. 2007;293:L870–82. [PubMed]
157. Chao PH, Lu HH, Hung CT, Nicoll SB, Bulinski JC. Effects of applied DC electric field on ligament fibroblast migration and wound healing. Connect Tissue Res. 2007;48:188–97. [PubMed]
158. Raja, Sivamani K, Garcia MS, Isseroff RR. Wound re-epithelialization: modulating keratinocyte migration in wound healing. Front Biosci. 2007;12:2849–68. [PubMed]
159. Chifflet S, Hernandez JA, Grasso S. A possible role for membrane depolarization in epithelial wound healing. Am J Physiol Cell Physiol. 2005;288:C1420–30. [PubMed]
160. Chifflet S, Hernandez JA, Grasso S, Cirillo A. Nonspecific depolarization of the plasma membrane potential induces cytoskeletal modifications of bovine corneal endothelial cells in culture. Exp Cell Res. 2003;282:1–13. [PubMed]
161. Borgens RB, Toombs JP, Breur G, Widmer WR, Waters D, Harbath AM, et al. An imposed oscillating electrical field improves the recovery of function in neurologically complete paraplegic dogs. J Neurotrauma. 1999;16:639–57. [PubMed]
162. Borgens RB, Blight AR, McGinnis ME. Behavioral recovery induced by applied electric fields after spinal cord hemisection in guinea pig. Science. 1987;238:366–9. [PubMed]
163. Borgens RB, Roederer E, Cohen MJ. Enhanced spinal cord regeneration in lamprey by applied electric fields. Science. 1981;213:611–7. [PubMed]
164. Shapiro S, Borgens R, Pascuzzi R, Roos K, Groff M, Purvines S, et al. Oscillating field stimulation for complete spinal cord injury in humans: a phase 1 trial. J Neurosurg Spine. 2005;2:3–10. [PubMed]
165. Smith SD. Effects of electrical fields upon regeneration in the metazoa. American Zoology. 1970;10:133–40. [PubMed]
166. Borgens RB. Are limb development and limb regeneration both initiated by an integumentary wounding? A hypothesis Differentiation. 1984;28:87–93. [PubMed]
167. Borgens RB, McGinnis ME, Vanable JW, Jr, Miles ES. Stump currents in regenerating salamanders and newts. Journal of Experimental Zoology. 1984;231:249–56. [PubMed]
168. Jenkins LS, Duerstock BS, Borgens RB. Reduction of the current of injury leaving the amputation inhibits limb regeneration in the red spotted newt. Developmental Biology. 1996;178:251–62. [PubMed]
169. Hotary KB, Robinson KR. Evidence of a role for endogenous electrical fields in chick embryo development. Development. 1992;114:985–96. [PubMed]
170. Sisken BF. Electrical-Stimulation of Nerves and Their Regeneration. Bioelectrochem Bioenergetics. 1992;29:121–26.
171. Sisken BF, Walker J, Orgel M. Prospects on Clinical-Applications of Electrical-Stimulation for Nerve Regeneration. J Cell Biochem. 1993;51:404–09. [PubMed]
172. Sharma KK, Niazi IA. Restoration of limb regeneration ability in frog tadpoles by electrical stimulation. Indian Journal of Experimental Biology. 1990;28:733–38. [PubMed]
173. Smith SD. Induction of partial limb regeneration in Rana pipiens by galvanic stimulation. Anatomical Record. 1967;158:89. [PubMed]
174. Borgens RB, Vanable JW, Jr, Jaffe LF. Bioelectricity and regeneration. I. Initiation of frog limb regeneration by minute currents. Journal of Experimental Zoology. 1977;200:403–16. [PubMed]
175. Smith SD. The role of electrode position in the electrical induction of limb regeneration in subadult rats. Bioelectrochem Bioenergetics. 1981;8:661–70.
176. Sisken BF, Fowler I. Induction of Limb Regeneration in the Chick-Embryo. Anatomical Record. 1981;199:A238–A39.
177. Becker RO. Stimulation of partial limb regeneration in rats. Nature. 1972;235:109–11. [PubMed]
178. Sisken BF, Fowler I, Romm S. Response of amputated rat limbs to fetal nerve tissue implants and direct current. J Orthop Res. 1984;2:177–89. [PubMed]
179. Masuda CA, Montero-Lomeli M. An NH2-terminal deleted plasma membrane H+-ATPase is a dominant negative mutant and is sequestered in endoplasmic reticulum derived structures. Biochem Cell Biol. 2000;78:51–8. [PubMed]
180. Reshkin SJ, Bellizzi A, Albarani V, Guerra L, Tommasino M, Paradiso A, et al. Phosphoinositide 3-kinase is involved in the tumor-specific activation of human breast cancer cell Na(+)/H(+) exchange, motility, and invasion induced by serum deprivation. J Biol Chem. 2000;275:5361–9. [PubMed]
181. Fukumoto T, Kema IP, Levin M. Serotonin signaling is a very early step in patterning of the left-right axis in chick and frog embryos. Curr Biol. 2005;15:794–803. [PubMed]
182. Denker SP, Barber DL. Cell migration requires both ion translocation and cytoskeletal anchoring by the Na-H exchanger NHE1. J Cell Biol. 2002;159:1087–96. [PMC free article] [PubMed]
183. Denker SP, Barber DL. Ion transport proteins anchor and regulate the cytoskeleton. Curr Opin Cell Biol. 2002;14:214–20. [PubMed]
184. Jaffe LF. Localization in the developing Fucus egg and the general role of localizing currents. Adv Morphog. 1968;7:295–328. [PubMed]
185. Kalmijn AJ. Electric and magnetic field detection in elasmobranch fishes. Science. 1982;218:916–8. [PubMed]
186. Kalmijn AJ. The electric sense of sharks and rays. J Exp Biol. 1971;55:371–83. [PubMed]
187. Gruler H, Nuccitelli R. Neural Crest Cell Galvanotaxis - New Data and a Novel-Approach to the Analysis of Both Galvanotaxis and Chemotaxis. Cell Motil Cytoskeleton. 1991;19:121–33. [PubMed]
188. Sasaki M, Gonzalez-Zulueta M, Huang H, Herring WJ, Ahn S, Ginty DD, et al. Dynamic regulation of neuronal NO synthase transcription by calcium influx through a CREB family transcription factor-dependent mechanism. Proc Natl Acad Sci U S A. 2000;97:8617–22. [PubMed]
189. Malhi H, Irani AN, Rajvanshi P, Suadicani SO, Spray DC, McDonald TV, et al. KATP channels regulate mitogenically induced proliferation in primary rat hepatocytes and human liver cell lines. Implications for liver growth control and potential therapeutic targeting. J Biol Chem. 2000;275:26050–7. [PubMed]
190. Levin M, Buznikov GA, Lauder JM. Of minds and embryos: left-right asymmetry and the serotonergic controls of pre-neural morphogenesis. Dev Neurosci. 2006;28:171–85. [PubMed]
191. Arcangeli A, Becchetti A. Complex functional interaction between integrin receptors and ion channels. Trends Cell Biol. 2006;16:631–9. [PubMed]
192. Cherubini A, Hofmann G, Pillozzi S, Guasti L, Crociani O, Cilia E, et al. Human ether-ago-go-related gene 1 channels are physically linked to beta1 integrins and modulate adhesion-dependent signaling. Mol Biol Cell. 2005;16:2972–83. [PMC free article] [PubMed]
193. Hegle AP, Marble DD, Wilson GF. A voltage-driven switch for ion-independent signaling by ether-a-go-go K+ channels. Proc Natl Acad Sci U S A. 2006;103:2886–91. [PubMed]
194. Mazzanti M, Bustamante JO, Oberleithner H. Electrical dimension of the nuclear envelope. Physiol Rev. 2001;81:1–19. [PubMed]
195. Tao Y, Yan D, Yang Q, Zeng R, Wang Y. Low K+ promotes NF-kappaB/DNA binding in neuronal apoptosis induced by K+ loss. Mol Cell Biol. 2006;26:1038–50. [PMC free article] [PubMed]
196. Yang SJ, Liang HL, Ning G, Wong-Riley MT. Ultrastructural study of depolarization-induced translocation of NRF-2 transcription factor in cultured rat visual cortical neurons. Eur J Neurosci. 2004;19:1153–62. [PubMed]
197. Murata Y, Iwasaki H, Sasaki M, Inaba K, Okamura Y. Phosphoinositide phosphatase activity coupled to an intrinsic voltage sensor. Nature. 2005;435:1239–43. [PubMed]
198. Li L, Liu F, Salmonsen RA, Turner TK, Litofsky NS, Di Cristofano A, et al. PTEN in neural precursor cells: regulation of migration, apoptosis, and proliferation. Mol Cell Neurosci. 2002;20:21–9. [PubMed]
199. Zhang W, Kone BC. NF-kappaB inhibits transcription of the H(+)-K(+)-ATPase alpha(2)-subunit gene: role of histone deacetylases. Am J Physiol Renal Physiol. 2002;283:F904–11. [PubMed]
200. Jouhou H, Yamamoto K, Homma A, Hara M, Kaneko A, Yamada M. Depolarization of isolated horizontal cells of fish acidifies their immediate surrounding by activating V-ATPase. J Physiol. 2007;585:401–12. [PubMed]
201. Morokuma J, Blackiston D, Levin M. KCNQ1 and KCNE1 K+ channel components are involved in early left-right patterning in Xenopus laevis embryos. Cell Physiol Biochem. 2008;21:357–72. [PMC free article] [PubMed]
202. Aw S, Adams DS, Qiu D, Levin MH. K-ATPase protein localization and Kir4.1 function reveal concordance of three axes during early determination of left-right asymmetry. Mech Dev. 2008;125:353–72. [PMC free article] [PubMed]
203. Shimeld SM, Levin M. Evidence for the regulation of left-right asymmetry in Ciona intestinalis by ion flux. Dev Dyn. 2006;235:1543–53. [PubMed]
204. Gillespie JI. The distribution of small ions during the early development of Xenopus laevis and Ambystoma mexicanum embryos. J Physiol. 1983;344:359–77. [PubMed]
205. Fang KS, Ionides E, Oster G, Nuccitelli R, Isseroff RR. Epidermal growth factor receptor relocalization and kinase activity are necessary for directional migration of keratinocytes in DC electric fields. J Cell Sci. 1999;112:1967–78. [PubMed]
206. Poo MM, Robinson KR. Electrophoresis of Concanavalin-a Receptors Along Embryonic Muscle-Cell Membrane. Nature. 1977;265:602–05. [PubMed]
207. Hotary KB, Robinson KR. Endogenous electrical currents and the resultant voltage gradients in the chick embryo. Dev Biol. 1990;140:149–60. [PubMed]
208. Borgens RB, Shi R. Uncoupling histogenesis from morphogenesis in the vertebrate embryo by collapse of the transneural tube potential. Developmental Dynamics. 1995;203:456–67. [PubMed]
209. Esser AT, Smith KC, Weaver JC, Levin M. Mathematical model of morphogen electrophoresis through gap junctions. Dev Dyn. 2006;235:2144–59. [PubMed]
210. Burr HS. Changes in the field properties of mice with transplanted tumors. Yale Journal of Biology & Medicine. 1941;13:783–88. [PMC free article] [PubMed]
211. Kauffman SA. The origins of order: self-organization and selection in evolution. xviii. Oxford University Press; New York: 1993. p. 709.
212. Martens JR, O’Connell K, Tamkun M. Targeting of ion channels to membrane microdomains: localization of KV channels to lipid rafts. Trends Pharmacol Sci. 2004;25:16–21. [PubMed]
213. Davies A, Douglas L, Hendrich J, Wratten J, Tran Van Minh A, Foucault I, et al. The calcium channel alpha2delta-2 subunit partitions with CaV2.1 into lipid rafts in cerebellum: implications for localization and function. J Neurosci. 2006;26:8748–57. [PubMed]
214. Wallace R. Neural membrane microdomains as computational systems: Toward molecular modeling in the study of neural disease. Biosystems. 2007;87:20–30. [PubMed]
215. Kline D, Robinson KR, Nuccitelli R. Ion currents and membrane domains in the cleaving Xenopus egg. J Cell Biol. 1983;97:1753–61. [PMC free article] [PubMed]
216. Perona R, Serrano R. Increased pH and tumorigenicity of fibroblasts expressing a yeast proton pump. Nature. 1988;334:438–40. [PubMed]
217. Megason SG, Fraser SE. Imaging in systems biology. Cell. 2007;130:784–95. [PubMed]
218. Adams DS, Levin M. In: Analysis of Growth Factor Signaling in Embryos. Whitman M, Sater AK, editors. Taylor and Francis Books; 2006. pp. 177–262.
219. Smith PJS, Sanger RS, Messerli MA. In: Methods and New Frontiers in Neuroscience. Michael AC, editor. CRC Press; 2007. pp. 373–405.
220. Reid B, Nuccitelli R, Zhao M. Non-invasive measurement of bioelectric currents with a vibrating probe. Nature protocols. 2007;2:661–9. [PubMed]
221. Moschou EA, Chaniotakis NA. Ion-partitioning membrane-based electrochemical sensors. Anal Chem. 2000;72:1835–42. [PubMed]
222. Steinberg BE, Touret N, Vargas-Caballero M, Grinstein S. In situ measurement of the electrical potential across the phagosomal membrane using FRET and its contribution to the proton-motive force. Proc Natl Acad Sci U S A. 2007;104:9523–8. [PubMed]
223. Yun Z, Zhengtao D, Jiachang Y, Fangqiong T, Qun W. Using cadmium telluride quantum dots as a proton flux sensor and applying to detect H9 avian influenza virus. Anal Biochem. 2007;364:122–7. [PubMed]
224. Wolff C, Fuks B, Chatelain P. Comparative study of membrane potential-sensitive fluorescent probes and their use in ion channel screening assays. J Biomol Screen. 2003;8:533–43. [PubMed]
225. Tyner KM, Kopelman R, Philbert MA. “Nanosized voltmeter” enables cellular-wide electric field mapping. Biophys J. 2007;93:1163–74. [PubMed]
226. Yu K, Ruan DY, Ge SY. Three electrophysiological phenotypes of cultured human umbilical vein endothelial cells. Gen Physiol Biophys. 2002;21:315–26. [PubMed]
227. Adams DS, Levin M. Inverse drug screens: a rapid and inexpensive method for implicating molecular targets. Genesis. 2006;44:530–40. [PMC free article] [PubMed]
228. Levin M. Is the early left-right axis like a plant, a kidney, or a neuron? The integration of physiological signals in embryonic asymmetry. Birth Defects Res C Embryo Today. 2006;78:191–223. [PubMed]
229. Adams DS. A new tool for tissue engineers: ions as regulators of morphogenesis during development and regeneration. Tissue engineering. 2008;14:1461–8. [PubMed]
230. Fischbarg J, Diecke FP. A mathematical model of electrolyte and fluid transport across corneal endothelium. J Membr Biol. 2005;203:41–56. [PubMed]
231. Pandiella A, Magni M, Lovisolo D, Meldolesi J. The effect of epidermal growth factor on membrane potential. Rapid hyperpolarization followed by persistent fluctuations. J Biol Chem. 1989;264:12914–21. [PubMed]
232. Lang F, Friedrich F, Kahn E, Woll E, Hammerer M, Waldegger S, et al. Bradykinin-induced oscillations of cell membrane potential in cells expressing the Ha-ras oncogene. J Biol Chem. 1991;266:4938–42. [PubMed]
233. Hulser DF, Lauterwasser U. Membrane potential oscillations in homokaryons. An endogenous signal for detecting intercellular communication. Exp Cell Res. 1982;139:63–70. [PubMed]
234. Dryselius S, Lund PE, Gylfe E, Hellman B. Variations in ATP-sensitive K+ channel activity provide evidence for inherent metabolic oscillations in pancreatic beta-cells. Biochem Biophys Res Commun. 1994;205:880–5. [PubMed]
235. Grapengiesser E, Berts A, Saha S, Lund PE, Gylfe E, Hellman B. Dual effects of Na/K pump inhibition on cytoplasmic Ca2+ oscillations in pancreatic beta-cells. Arch Biochem Biophys. 1993;300:372–7. [PubMed]
236. Stokes CL, Rinzel J. Diffusion of extracellular K+ can synchronize bursting oscillations in a model islet of Langerhans. Biophys J. 1993;65:597–607. [PubMed]
237. Aeckerle S, Wurster B, Malchow D. Oscillations and cyclic AMP-induced changes of the K+ concentration in Dictyostelium discoideum. Embo J. 1985;4:39–43. [PubMed]
238. Giulian D, Diacumakos EG. The electrophysiological mapping of compartments within a mammalian cell. J Cell Biol. 1977;72:86–103. [PMC free article] [PubMed]
239. Liebau S, Propper C, Bockers T, Lehmann-Horn F, Storch A, Grissmer S, et al. Selective blockage of Kv1.3 and Kv3.1 channels increases neural progenitor cell proliferation. J Neurochem. 2006;99:426–37. [PubMed]
240. Banghart M, Borges K, Isacoff E, Trauner D, Kramer RH. Light-activated ion channels for remote control of neuronal firing. Nat Neurosci. 2004;7:1381–6. [PMC free article] [PubMed]
241. Chambers JJ, Banghart MR, Trauner D, Kramer RH. Light-induced depolarization of neurons using a modified Shaker K(+) channel and a molecular photoswitch. J Neurophysiol. 2006;96:2792–6. [PubMed]
242. Gradinaru V, Thompson KR, Deisseroth K. eNpHR: a Natronomonas halorhodopsin enhanced for optogenetic applications. Brain cell biology. 2008;36:129–39. [PMC free article] [PubMed]
243. Miesenbock G, De Angelis DA, Rothman JE. Visualizing secretion and synaptic transmission with pH-sensitive green fluorescent proteins. Nature. 1998;394:192–5. [PubMed]
244. Tsutsui H, Karasawa S, Okamura Y, Miyawaki A. Improving membrane voltage measurements using FRET with new fluorescent proteins. Nat Methods. 2008;5:683–5. [PubMed]
245. Zhao S, Cunha C, Zhang F, Liu Q, Gloss B, Deisseroth K, et al. Improved expression of halorhodopsin for light-induced silencing of neuronal activity. Brain cell biology. 2008;36:141–54. [PMC free article] [PubMed]
246. Zhang F, Wang LP, Brauner M, Liewald JF, Kay K, Watzke N, et al. Multimodal fast optical interrogation of neural circuitry. Nature. 2007;446:633–9. [PubMed]
247. Wang H, Peca J, Matsuzaki M, Matsuzaki K, Noguchi J, Qiu L, et al. High-speed mapping of synaptic connectivity using photostimulation in Channelrhodopsin-2 transgenic mice. Proc Natl Acad Sci U S A. 2007;104:8143–8. [PubMed]
248. Stern C. Experimental reversal of polarity in chick embryo epiblast sheets in vitro. Exp Cell Res. 1982;140:468–71. [PubMed]
249. Woodruff RI. Calmodulin transit via gap junctions is reduced in the absence of an electric field. J Insect Physiol. 2005;51:843–52. [PubMed]
250. Kurtz I, Schrank AR. Bioelectrical properties of intact and regenerating earthworms Eisenia foetida. Physiol Zool. 1955;28:322–30.
251. Uzman JA, Patil S, Uzgare AR, Sater AK. The role of intracellular alkalinization in the establishment of anterior neural fate in Xenopus. Developmental Biology. 1998;193:10–20. [PubMed]