David W. Sretavan et al- Microscale Surgery on Single Axons

8/3/2019 David W. Sretavan et al- Microscale Surgery on Single Axons

http://slidepdf.com/reader/full/david-w-sretavan-et-al-microscale-surgery-on-single-axons 1/12



require up to 1 to 2 years to grow back to their target cells aftersevere injury. During this extended period, neurons can diefrom the lack of retrograde trophic support, and denervatedtarget tissues such as muscles can atrophy. All of these factorslead to the fact that although recovery occurs, full recoveryafter significant peripheral nerve injury is the exception rather

than the rule.

CURRENT STRATEGIES FOR CENTRALNERVOUS SYSTEM AXON REGENERATION

Although PNS axon regeneration is possible under favor-able circumstances, there is no meaningful axon regenerationin the central nervous system (CNS). The lack of CNS axonregeneration is thought to be in part the result of inhibitoryproteins associated with adult CNS myelin (2, 11, 16), inhibi-tory proteoglycans upregulated at the site of CNS injury (32),and an intrinsic downregulation of the growth ability of adultneurons (12, 22). Current research and therapeutic strategiesinclude the promotion of axon regeneration from the site ofdamage back to original neuronal targets (25, 47, 49) and thetension-mediated stretching of axons to bridge regions ofnerve injury (50). In approaches using regeneration strategies,axons not only must be coaxed to grow over long distances, but also this growth must be directed back to specific neuronaltargets and not form abnormal neuronal circuitry. Further-more, regenerating axons must somehow reestablish the orig-inal highly specific pattern of synaptic contacts on individualpostsynaptic cells. Last, all of this must occur rapidly enough before denervation atrophy of postsynaptic neurons (15, 24,51) or apoptosis of the original damaged neurons (1).

SURGICAL REPAIR OF AXONS:VENTURING INTO THE MICROSCALE

As an alternative to axon regeneration, we propose a novelnerve repair paradigm in which the damaged region of indi-vidual axons is excised and replaced by healthy donor axonsegments to reestablish neuronal connectivity and function(Fig 1A). Direct axon repair can be broken down into threeessential steps, beginning first with axon cutting to excisedamaged axon regions, leaving clean healthy axon ends. Next,a donor axon segment is brought in to fill the gap between theends of the host axons. As soon as the ends of the host anddonor axons are aligned and apposed, the axon segments thenare fused to establish functional integrity. A similar sequenceof axon alignment and fusion also would apply in the instanceof injuries in which nerve mobilization can eliminate gapsresulting from damage, and thus eliminate the need for donoraxon segments (Fig. 1B). Compared with strategies based onaxon regeneration, we propose that axon repair should beundertaken soon after nerve injury, before Wallerian degen-eration and the loss of the distal axon segment with its syn-aptic connections. If successful, the reestablishment of axonalcontinuity, together with the presence of an intact synaptic

branching pattern, could lead to recovery of neuronal func-tion.

To our knowledge, a method for the systematic and con-trolled repair of individual axons has not been proposed pre-viously. The manipulation of axons whose diameter may be inthe micron to submicron range is not feasible using currentmicrosurgical tools and techniques. However, it is precisely atthese small-length scales that microtechnology and nanotech-nology excel. The potential impact of microtechnology and

nanotechnology on the neurosurgical operative environmenthas been recognized, and these emerging fields have beenpredicted to form the scientific foundations of new paradigms(31, 44). With these ideas in mind, our multidisciplinary re-search group set out 4 years ago to obtain proof of principlefor the basic steps of axon repair. In this article, we discuss anddemonstrate some of the microscale core technologies we believe are highly promising for axon repair. In addition, wealso present our prototype miniature multifunctional axonsurgery platform that is approximately 1 mm3 and that con-tains some of the functionality necessary for axon surgery. Byexploiting advantages of microfabrication, noncontact electro-kinetic methods for axon manipulation, and fundamental

principles of neurobiology, it may be feasible in the future todevelop Micro Electro-Mechanical Systems (MEMS) surgicalmicrodevices that will allow neurosurgeons to engage in di-rect axon repair at a cellular level.

MICROFABRICATED DEVICES FOR AXONCUTTING

We have used MEMS microengineering fabrication tech-niques to develop small surgical devices with nanoscale fea-

FIGURE 1. Diagram showing the proposed scheme for axon repair. A, the gray region in the middle of the axon represents the injured segment. InStep 1, the damaged axon region is excised. In Step 2, a segment of healthy donor axon is brought into place. In Step 3, the host and donoraxons are joined together to reestablish functional continuity. B, ininstances where a damaged nerve can be mobilized to eliminate a gap

between the distal and proximal ends, a similar sequence of axon align-ment and joining can be used without resorting to the use of donor axons.In Step 1, the damaged region is excised. In Step 2, the nerve is mobilizedand the axons are aligned. In Step 3, the axons are joined.

SRETAVAN ET AL.

636 | VOLUME 57 | NUMBER 4 | OCTOBER 2005 www.neurosurgery-online.com



tures capable of repeatable, precise cutting of single axons.MEMS has its roots in the semiconductor industry in whichlithography, chemical etching, and silicon doping are used inthe manufacturing of integrated circuits. The ability of theseprocessing methods to create precise microscale features insilicon was adopted for the manufacture of micron-sized de-

vices. Rather than the electronic properties of single crystalsilicon, MEMS exploits silicon’s excellent mechanical proper-ties (40). With the addition of sensors, force generating actu-ation mechanisms, as well as electronic controls, MEMS mi-crodevices are capable of carrying out useful function atunprecedented microscale levels. Given that the mechanicalstrength of silicon is superior to that of steel (48), MEMSdevices are robust and have been used in everyday productssuch as automobile accelerometers, ink jet printer heads, andvideo projectors. An active area of MEMS research is thedevelopment of biomedical microdevices for diagnostics andhealthcare (42, 44). BioMEMS devices are well suited for han-dling the small tissue or fluid samples often typical of biotech-

nology research and medical diagnostics. The small size ofBioMEMS devices also opens new opportunities for usewithin the human body for imaging, cellular diagnosis, andtissue engineering.

DESIGN OF CUTTING DEVICE

Our current cutting device consists of a silicon nitride knifewith an ultrasharp knife edge (Fig. 2A) mounted onto a silicon-

based compliant knife suspension (Fig. 2, B and C). The knifeedge has a radius of curvature of only approximately 20 nm,similar to the diameter of a single microtubule (32) or thewidth of synaptic clefts (29). The knife is constructed from athin silicon nitride membrane and is effectively transparent,allowing the optical monitoring of axons during the cutting

procedure (Fig. 2D). The mechanical compliance of the sus-pension can be tuned to any desired stiffness within a range todeliver sufficient force for cutting various biological tissuesfrom single axons to the harvesting of specific cell populationsfrom histological tissue sections.

CUTTING DEVICE PERFORMANCE

These cutting devices are capable of reliable and preciseaxon cutting (Fig. 3) that leave clean edges and are superior tocurrent cell harvesting methods using razor blades or glassmicropipettes that tend to shear and tear axons. Althoughsuch imprecise shearing may be acceptable in tissue harvest-

ing for cellular or biochemical studies, these methods cannot

FIGURE 2. Photographs showing the microfabricated axon cuttingdevice. A, axon knife assembled within a compliant suspension frame. The

pyramidal structure in the center is an optically transparent axon knifewith a 10-m cutting edge. f, silicon suspension flexures; k, axon knife; h,handle to micromanipulator. The footprint of the frame is 1 mm2. Scale 200 m. Devices with substantially smaller footprints can be fabricated. B,detail of the silicon suspension flexure on each side of the axon knife that

provides mechanical compliance during cutting. The number of switch-backs in the flexure can be modified to obtain devices with differentmechanical compliances that deliver a range of cutting forces. The compli-ance of the flexures allows it to act as a suspension, maximizing the dura-bility of the knife edge. C, oblique view of the silicon-nitride knife. Kniveswith edges from 5 to 200 m have been fabricated and tested. A 200-mlong knife is shown. D, scanning electron micrograph showing a cross sec-tion of the silicon-nitride knife at its very edge. The radius of curvature atthe edge is roughly 20 nm. Scale 100 nm.

FIGURE 3. Images demonstrating an axon cutting sequence and examples of cut axons. A, axon to be cut (arrow) in center of field. Knife is attop (not in the plane of focus). B, knife positioned over axon by a micro-manipulator. Note that the axon is visible through the knife. C, knife islowered for cutting. D, the resulting cut axon. E, another axon before cut-ting. F, same axon as in (E) after cutting. G, examples of myelinated(top) and unmyelinated ( bottom) adult mouse sciatic nerve axons thathave been cut. D–G, scale 10 m.

MICROSCALE AXON SURGERY

NEUROSURGERY VOLUME 57 | NUMBER 4 | OCTOBER 2005 | 637



be used for axon repair. Laser thermal ablation can performprecise cutting (6) but typically involves large instrumentationthat is difficult to integrate with other functional componentsrequired for repair at the micron scale of axons. Moreover,laser cutting typically is achieved by the thermal vaporizationof tissue within a minimum spot size of roughly 0.5 m in

diameter.Both adult PNS (sciatic nerve) and CNS (optic nerve) axons

from mice rapidly reseal after they have been severed usingthe microcutting devices. Invertebrate axons and mammalianembryonic axons have a self-repair resealing mechanism afterinjury involving calcium-mediated exocytosis (14, 21, 23, 53,61). Based on studies designed to detect the leakage of solublecytoplasmic green fluorescent protein (GFP) from axons aftercutting, we have found that this resealing ability also is evi-dent in adult axons. This finding suggests that axons toleratecutting by a microfabricated ultrasharp microknife. The pro-tein and cell signaling cascades necessary for physiologicalself-resealing at the cut ends is operational and the intact axon

segments can be used in the subsequent steps of axon repair.Future improvements to the microcutting device include

sensors as well as force-generating actuation mechanisms thatautomatically will deliver a controlled cutting stroke. Bothpiezoelectric and thermal expansion actuation mechanismscan deliver forces in the range needed for axon cutting. Amicrocutting device with on-board sensing and actuation canfunction as a semiautonomous instrument, requiring only ini-tiating commands from the human operator. The eliminationof any need for manual manipulation maximally uses theprecision of MEMS microdevices.

DIELECTROPHORESIS FOR AXONMANIPULATION

After cutting, the host and donor axon segments must bealigned with close membrane apposition in preparation forsubsequent fusion. We have chosen dielectrophoresis (DEP) asthe method of choice for noncontact axon manipulation at themicroscale. DEP is the movement of small polarizable objectssuch as cells in a nonhomogeneous AC electrical field and has been used effectively as a electrokinetic method to move andsort cells within microdevices (19, 55). In essence, objects in analternating electrical field experience field-induced polariza-tions on their surfaces. In the presence of a homogeneous field,dipole forces alternate on an object’s surface along with alter-nations in the electrical field, but no net movement of theobject occurs. However, in a nonhomogeneous field with gra-dients of field strengths, the object experiences a net force inone direction, causing it to move toward one side of theelectrical field (for review, see (20)). DEP is fundamentallydistinct from electrophoresis in which the net charges onobjects lead to movement in one direction within a stationaryelectrical field.

We have found that cylindrical structures such as axons can be manipulated in DEP fields generated by micron-sized elec-

trodes (Fig. 4). In this example, application of a DEP fieldcaused the axons to be deflected upward from the horizontalelectrode toward the top half of the semicircular electrode (Fig.4B). Axons returned to their original location after the stimu-lus was turned off (Fig. 4C). The direction, magnitude, andspeed of axon movement within the DEP field were main-

tained over several cycles of DEP field initiation. This partic-ular electrode configuration moved axons at roughly 5 m/sec. In ongoing experiments, we are testing various electrodeconfigurations that reliably will bring together and align theends of two separate axons as required for the proposedmethod of axon repair outlined in Figure 1. The viability ofcells that have undergone DEP has been studied in somedetail. Fibroblasts and erythroleukemia cells show good viability as determined by trypan blue dye exclusion and contin-ued cell proliferation after DEP (18, 56). In recent work, cor-tical neurons that have been subjected to DEP are capable ofaxon outgrowth (26, 27), indicating that axon physiology is notaffected by DEP and is directly relevant to our proposed use of

DEP for axon manipulation.

DEP ELECTRODE ARRAYS

The effective DEP fields for axon manipulation are rela-tively short ranged and extend only approximately 30 maway from the electrodes. This limited range can be veryuseful for isolating peak DEP forces to specific axons of inter-est without much effect on neighboring axons in the vicinity.In addition, it should be possible to use closely spaced elec-

FIGURE 4. Photomicrographsdemonstrating noncontact DEPmanipulation of axons. Adult seg-ments of sciatic nerve were har-vested and digested with an enzymecocktail to remove the ensheathingtissue and to release individual ax-ons. The DEPelectrodes consist of a

pair of 10-m wide metal traces( black) deposited onto glass. A, ar-rows point to axons to be subjectedto DEP fields. B, after activation of DEP field, axons moved away from the horizontal electrode toward the semicir-cular electrode. C, axons returned to original position after termination of DEP

field. The axons moved 10 m in 1 to 2 seconds. Scale 20 m.

SRETAVAN ET AL.




trode arrays (Fig. 5) to move axons at will to desired locationswithin a surgical field. The sequential activation of a series ofaddressable electrodes within the array should generate prop-agating waves of DEP forces capable of directing the move-ment and alignment of axons. Because DEP fields are deter-mined by electrical parameters and its theoretical basis is

relatively well understood, this method of axon manipulationcan be conveniently preprogrammed. We envision that giventhe starting location of an axon of interest and a sufficientunderstanding of axon behavior in DEP fields, it should bepossible to obtain a computer-generated set of DEP parame-ters and electrode activation sequences that will move theaxon to any desired position. Computer-controlled activationsequences then can be executed on human command to trig-ger optimal stimulus parameters for axon manipulation andalignment.

AXON ELECTROFUSION

After axon cutting and alignment, the third step of axonrepair is the joining of axon ends to achieve functional conti-nuity. Cell fusion in biology is most commonly used in thecreation of hybridoma cells for monoclonal antibody produc-tion. A number of methods can be used to promote cell fusion.These include chemical fusion using polyethylene glycol,laser-induced fusion, and electrofusion. We have selected elec-trofusion as our initial technology for axon fusion. Electrofu-sion results in the transient breakdown of cell membranes andthe formation of pores that are unstable and eventually reseal(38). If the pores are formed in the region of contact betweenneighboring cells, fusion between the two cells will occur. Thethreshold voltage across a membrane required for this electri-

cal breakdown is approximately 1 V. In cell culture, the typicalfield strengths required to generate 1 V across membranes arein the range of 2 to 8 kV/cm (5, 38), usually delivered usingtwo to four rectangular pulses of 10 to 100 s each.

Electrofusion has been used extensively to create mamma-lian cell hybrids. Its ease of use and fusion yield of two ordersof magnitude better than chemical polyethylene glycol fusion(28, 30) has made electrofusion frequently the technique ofchoice for hybridoma production. Electrofusion also has beenused to fuse embryonic or fetal somatic cells with enucleated

oocytes successfully to clone both sheep and mice (54, 57). Theability of fused embryo cells to develop to term, and subse-quently to grow into adult animals, argue for the safety andefficacy of electrofusion.

We favor the use of electrofusion for axon surgery. With theidentification of appropriate parameters, electrofusion can bepreprogrammed to automate axon fusion. In addition, becausewe will use DEP to align axons, electrodes already will beconveniently placed and electrical stimulation parameters can be switched conveniently from DEP to electrofusion. In axonrepair, the pairing of DEP with electrofusion permits bothsteps in the repair sequence to be accomplished using elec-trodes, simplifying the design and operation of the MEMS

axon surgery platform.

DEMONSTRATION OF AXON FUSION

The successful splicing together of two axons should createcytoplasmic continuity and the flow of protein components between the two axon segments. A direct observation of flowis feasible using axons from transgenic mice in which the geneencoding GFP has been introduced into the germ line, andGFP protein is produced in all the cells of the animal (39). GFPexists as a soluble protein in cells, causing cells to exhibit greenfluorescence on appropriate excitation (excitation

max, 489

nm; emission max

, 508 nm). Axons from nontransgenic wild-

type mice do not exhibit fluorescence under the same wave-length excitation.We have demonstrated successful fusion between GFP-

containing and non–GFP-containing axons leading to thespread of GFP between the fusion partners. These studieswere performed in culture on pairs of axons that cross and lieon top of each other (Fig. 6). Figure 6A shows two bundles ofretinal axons running from left to right in the field of study.The corresponding bright field picture (Fig. 6B) indicates thepresence of numerous other nonGFP retinal axons in the field,including the axon of interest indicated by the arrow. Electri-cal field stimulation was applied locally within 20 m of thetarget axon pair using a pair of 12-m diameter platinumelectrodes spaced 12 m apart. The appearance of GFP fluo-rescence in this retinal axon was observed within 5 minutesafter electrofusion, and the entire axon extending between thetwo axon bundles exhibited GFP fluorescence by 20 minutesafter electrofusion (Fig. 6C). The bright field appearance ofaxons 20 minutes after electrofusion is shown in Figure 6D. Asecond example of the appearance of GFP after electrofusionin a previously unlabeled axon is shown in Figure 6, E–H .Although these data are promising, additional work must becarried out to verify the central tenet of our proposed axonrepair scheme that axons that have been repaired are capable

FIGURE 5. Photomicrograph demonstrating an array of microelectrodes fabricated on a glass substrate by isotropic, wet etching and coated with athin transparent film of conductive indium tin oxide. When energized, thehigh gradients of electric field at the sharp electrode tips are designed to

generate DEP forces to attract axons to that location. Further develop-ments are underway to allow each electrode to be addressed individually sothat axons can be directed to specific locations within the array. Scale 100 m.





of normal function and survival. Aspects of axonal function ofinterest include the ability for electrical conduction and themaintenance of axonal transport.

ASSEMBLING CORE TECHNOLOGIES INTOA MULTIFUNCTIONAL AXON REPAIRMICRODEVICE

The results presented thus far were obtained using earlyprototypes of MEMS axon knives and electrodes, thus under-scoring the robustness and ease of use of the underlyingprinciples. However, to develop a practical methodology foraxon repair, the individual axon repair steps have to be coor-dinated into an efficient sequence that is applicable at themicroscale of axons. Our strategy is to provide all of thefunctions necessary for axon repair within a multifunctionalMEMS axon surgery platform. Although it may be technicallyfeasible to perform each step involved in axon repair by op-erating multiple microscale tools, each independentlymounted on its own positioner (similar to probe stations fortesting integrated circuits), such an arrangement would beextremely bulky, inefficient, and time consuming to use clin-ically. Our approach solves these problems by assembling allrepair functions onto a single MEMS surgical platform thatincorporates as much autonomy as possible into each repairstep. Such a semiautonomous microscale surgical platformwill achieve the degree of miniaturization and performanceefficiency required for device use at a cellular level.

To demonstrate this concept,we designed and fabricated aprototype of an integrated axonsurgery platform that is 1 mm3 insize (Fig. 7). This prototype uses aspace frame (Fig. 7A) in which

interlocking and latching featuresare used for self-fixation and tofacilitate assembly. Individualcomponents needed for axon re-pair such as the microcutting de-vice and DEP/fusion electrodearrays are inserted and attachedto the space frame. With the baseof the various components fixedwithin the space frame, their rel-evant functional elements such asthe axon knife and electrodes allextend toward the bottom sur-

face of the cubic device closest tothe target axons, where they arepositioned in an optimal spatialrelationship with one another toexecute axon repair (Fig. 7, B–D).This modular design also allowscomponents to be switched con-veniently in and out during ex-

perimentation and with design improvements. In addition, theopen structure allows for optical monitoring of the repair pro-cess. Features such as microfluidics to control the fluid environ-ment or to deliver reagents to assist axon repair can be readilyincorporated into the platform. Last, a cubic framework can

incorporate on-board actuation mechanisms to move the knife inan up-and-down motion, allowing the autonomous execution ofcutting on human command, and maximally can benefit from theprecision offered by microscale devices.

An example of an axon surgery platform prototype consist-ing of an assembled space frame with a microcutting device isshown in Figure 7, E and F. This 1-mm3 prototype platformwas assembled using customized microfabricated microgrip-pers and microtools specifically designed for micron-scaleassembly. With existing microfabrication technologies, itshould be quite possible to develop improved versions of axonsurgery platforms that are substantially smaller in size. Al-though the current cubic design offers many advantages, thisdesign is by no means fixed, and eventual multifunctionaldevices capable of the full range of axon repair steps may takeon quite different shapes.

INTEGRATING MICROSCALE AXONSURGERY PLATFORMS INTO THE

SURGICAL FIELD

To use any microscale device for surgery effectively, meth-ods must be developed to integrate such small devices for use

FIGURE 6. Examples of axon electrofusion resulting in spread of green fluorescent protein between retinalaxons in culture. A, two bundles of GFP axons in the field of study. B, bright field picture of the field shown in(A) indicating an unlabeled axon (arrow). C, appearance of GFP in the same axon (arrow) 20 minutes afterelectrofusion. D, bright field picture 20 minutes after electrofusion. Arrow indicates same axon as in C. E, GFP-labeled axons. F, bright field image of the same field. Arrow points to unlabeled axons. G, within 1 minute of electrofusion, GFP appeared in a previously unlabeled axon (arrow). H, arrow indicates same axon as in G 15minutes after fusion. A–H, scale 10 m.

SRETAVAN ET AL.




within a surgical field. One method is through the use of a

specialized surgical chamber such as the one shown in Figure8. Some features that are likely important for such chambersinclude a means of holding the host and donor nerves in place,maintenance of an appropriate fluid environment, and a me-chanical anchor and micromanipulator that can position thesmall axon surgery platform for repair. In Figure 8, the com-ponents within the chamber, including nerves, axons, and theaxon surgery platform, are designed to be mechanically iso-lated from the rest of the body (components are not drawn toscale). Such chambers could be positioned adjacent to periph-eral nerves relatively easily, where large open fields of surgi-cal exposure are used. A greater challenge would be to posi-tion such a device within the spinal cord or brain. Possible

solutions include the use of access ports and minimally inva-sive techniques, coupled with the enhanced dexterity madepossible by robotic devices. Together, these technologies con-ceivably could allow the placement and operation of a MEMSsurgical platform anywhere in the nervous system.

CLINICAL ISSUES

Currently, there is no specific treatment for axon damagefrom trauma either to the PNS or the CNS. Patients with

severe nerve injuries often are viewed as requiring chronicrehabilitative care, with those with CNS injuries as having aparticularly poor outlook. We think that with the advent of

microscale axon repair, the perception of nerve and axoninjuries may shift and may begin to be viewed as an acutesurgical situation. The ability to intervene and perform axonrepair before irreversible changes take place, such as Walle-rian degeneration, loss of synaptic contacts, or neuronal celldeath, will shift the emphasis from support and rehabilitationto earlier intervention and treatment. Given that our proposedmethod of axon repair is significantly different from currentpractice and takes place at an unprecedented small-lengthscale, a host of clinical issues must be considered and evalu-ated. Below, we lay out a number of these issues with the goalof stimulating discussion and further research into areasprompted by the possibility of neurosurgery at the microscale

level.

When Should Axon Repair Take Place?

An advantage of axon repair is that, if performed soon afterinjury, it may be possible to reuse the synaptic connections ofthe distal axon segment and to achieve rapid functional recov-ery. This consideration sets the onset of Wallerian degenera-tion as the upper limit for the interval from damage to repair.In the PNS, the interval between injury and onset of axonalchanges is 24 to 48 hours (8). In the CNS, Wallerian degener-ation occurs more slowly or to a lesser extent after axondamage (3, 34), and the interval after injury available forintervention to conduct axonal repair may be longer. Theexistence of a critical interval for axon repair will necessitateadjustments in clinical management in the immediate periodafter injury. In addition to recognition of nerve damage aspotentially requiring acute care, improved and faster methodsfor determining the severity of axon injury would be critical. Italso should be noted that basic neurobiological research is beginning to identify some of the cell signaling pathways inWallerian degeneration (60). It is possible that in the future,one may intervene to delay the onset of Wallerian degenera-tion and increase the window of opportunity for axon repair.

FIGURE 7. Diagrams showing the design and assembly of prototypemultifunctional axon surgery platform. A, schematic representation of the

space frame. Carrier holes are used for positioning the platform. B, modu-lar axon repair components such as cutting devices and electrode arraysare inserted into the space frame to preposition their functional elements

for efficient axon repair. (Generic components are shown here.) In currentconcept, the space frame also contains a force generation (actuation) mech-anism to execute the up–down motion of the axon knife during cutting. C,oblique view of platform assembled with a microcutting device and elec-trode arrays. D, view from bottom. E, 16 individual microfabricated partson display on a penny. A microgripper and a microprobe are shown on theleft. F, scanning electron microscope image of the assembled axon surgerydevice prototype containing a functioning axon microcutting device andsupporting pieces locked in place to form a 1 mm3 superstructure. Scale 200 m.

FIGURE 8. Conceptual drawing of a surgery chamber for axon repair.The surgery chamber provides mechanical support and environmental con-trol for the host and donor axons as well as the axon surgery platform.(Components are not drawn to scale). Such platforms also can be used incases where injured nerves are mobilized and proximal and distal ends of host axons are joined without the insertion of donor axons (see Fig. 1B).





Sources of Donor Axons

In current clinical management, procedures involving pe-ripheral nerve grafts are all performed at the nerve connectivetissue level, with the sole purpose of providing a connectivetissue conduit to encourage possible axon regeneration. Nerve

repair therefore is not aimed directly at repairing individualaxons. However, donor axons to be used as fusion partnersalso can be obtained from this type of autologous donor nervesegment (e.g., often the sural nerve in the lower leg). For axonrepair, the harvested nerve segments can provide a source ofhealthy donor axon segments to be used in axon fusion. Axonswill be separated from the surrounding connective tissues byenzymatic digestion of collagen and other means to eliminatethe Schwann cells in peripheral nerve (Fig. 9). The exposedaxons are then ready for cutting, alignment, and fusion.

How Many Axons Need to be Repaired?

An important point to consider is how many axons need to be repaired to obtain functional recovery. Results from studiesaiming to promote CNS axon regeneration can serve as aguide. After spinal cord lesions in rodents, the regeneration ofseveral hundred axons has been reported to correlate withsome recovery of function (10, 13, 37), and it is generallyagreed that the survival of a fraction of axons within a path-way is sufficient to maintain some degree of useful function (4,33, 43, 46, 59). This finding is thought to reflect the potentialfor substantial plasticity in the injured nervous system. If so,this indicates that it may not be necessary to repair all axonswithin a PNS nerve or a CNS axon tract. The immediate repairof a subset of axons may provide a sufficient degree of func-

tional recovery to alleviate the disability normally associatedwith nerve injuries. It should be recognized that only verycrude estimates of the required number of repaired axonscould be given at the present time. It seems likely that thenumber of axons to be repaired will depend on the site ofinjury, and that rational guidelines may be developed in thefuture based on direct clinical experience.

Which Axons to Repair?

In clinical applications, the careful selection of which axonsto repair will contribute significantly to surgical success. Be-cause the surgical repair must be completed before the onsetof distal Wallerian degeneration of axons, it will be possible to

electrophysiologically map and identify cut distal axons byelectrically stimulating them and recording from their targetstructures. One strategy is to focus on damaged axons that arephysically most accessible for MEMS-assisted repair. In addi-tion, it may be useful to target large-diameter axons for repair, because these axons mediate motor and somatosensory func-tion. Axon repair also may be accomplished more easily inlarger axons than in small axons that are less than 1 m indiameter. A further consideration is to identify situationswhere the reconnection of a subset of axons can make a largedifference in clinical management and in the patient’s qualityof life. An example is the restoration of partial movement inthe hand and digits, which would greatly impact how tet-raplegic patients interact with their environment. A secondexample is the repair of axon pathways mediating bladdercontrol. After spinal cord injury, loss of voluntary control ofvoiding leads to urinary retention and eventually to inconti-nence (58). Management of these conditions by catheterizationleads to urinary tract infections, which is a major source ofmorbidity in these patients (7). Recovery of some degree of bladder control would contribute significantly to their qualityof life. The development of microscale axon repair, matchedwith a well thought out surgical plan and realistic expecta-tions, will be important ingredients for clinical success.

Selecting Axon Fusion Partners

A key element in axon repair is the identification of appro-priate axon partners for reconnection. A characteristic featureof both CNS axon tracts and PNS nerves is the topographicorganization of axons. Throughout the nervous system, axonsthat originate from adjacent neurons tend to run together andto maintain their neighbor relationship within a nerve or axontract. For example, motor axons innervating a group of neighboring muscles typically are found next to each other in thesame subregion within an axon tract or nerve. Similarly, axonscarrying sensory information from adjacent areas of the skinsurface are found next to one another both in a peripheralnerve and in the spinal cord. During repair, it thus may not benecessary to use donor axon segments specifically to reconnectthe proximal axon segment with its original distal segment.Reconnection with an adjacent axon, as long as it is not anextreme mismatch, may restore acceptable function.

The plasticity and remodeling of neuronal circuitry that isknown to occur in the adult nervous system also may work infavor of axon repair. The extent to which this can occur isillustrated by the cortical plasticity that occurs in patients whohave received intercostal nerve to musculocutaneous nervetransfers and in whom biceps movement is gradually dissoci-ated from respiratory effort and eventually comes under vo-litional control (35, 36). Exactly how much of a mismatch will

FIGURE 9. Photomicrograph showing individual axons at the end of asegment of adult mouse sciatic nerve that was subjected to connective tis-sue digestion. A single myelinated axon is seen in the center of the field.Scale 20 m.

SRETAVAN ET AL.




be tolerated may depend on the specific part of the nervoussystem involved and the function of the axon populationunder consideration. Advances in new methodologies for di-rect axon repair we hope will encourage development of pre-operative or intraoperative methods of rapid functional map-ping of axon groups in both the PNS and CNS.

How Long Will It Take?

Axon repair will be a practical approach only if it can becarried out as a surgical procedure of reasonable duration.Our investigations of the core technologies and how quicklyeach repair step currently can be accomplished provide guid-ance for a minimum time required. As soon as positioned,actuated microdevices with reasonable force generation andgearing design should be able to deliver an axon cutting strokeconsisting of a 50-m length of travel in under 5 seconds. Wehave observed that axons move at approximately 5 m/s inDEP fields generated by very simple electrode configurations.If we estimate a need to move axons 25 to 50 m for properalignment with their fusion partner, this second step of repairshould take 5 to 10 seconds. Because the same electrodes will be used for the final fusion step, electrofusion can be initiatedimmediately after alignment. Typical electrofusion parametersare applied over 2 to 3 seconds. From these estimates, wearrive at roughly 20 seconds for each axon. If the protocol callsfor each of these steps to be completed on one axon at a time,180 cycles of axon repair can be performed in 1 hour.

A much more efficient way to use microtechnology, how-ever, is to exploit its potential for batch processing of function.In particular, the overall time required for axon repair can beshortened significantly by cutting and trimming multiple ax-ons simultaneously. In addition, DEP electrode arrays with

individually addressable electrodes will permit the simulta-neous manipulation of multiple axons within the surgicalfield. After alignment, the entire set of axons can then besubjected to electrofusion. Such a parallel processing approachwill increase significantly the number of axons that can berepaired per unit time and will make the repair of severalthousand axons a reasonable surgical goal.

Is Axon Repair Feasible in the CNS?

From a basic neurobiological perspective, axon repair meth-odologies for PNS axons also should be applicable to CNSaxons because of their fundamental similarities in biologicalstructure. Beyond basic neurobiology, however, access to theCNS may be more demanding and likely will require surgicalchambers of different design. A second issue is whether PNSaxons can be used as grafts to replace a segment of CNS axon.A definitive answer is not at hand because such an operationhas yet to be performed. It is well known, however, that CNSaxons will grow into a piece of PNS sciatic nerve graft forsignificant distances (1, 2). Although it is unclear whether andhow a CNS/PNS hybrid axon will function, the highly similarcell biological make-up of PNS and CNS axons suggests that itmay be worthwhile exploring this approach.

Remyelination of Repaired Axons

Schwann cells or oligodendrocyte glia cells extend cytoplas-mic protrusions to form myelin around axons to enhanceelectrical conduction. The importance of myelination is illus-trated by demyelinating diseases such as multiple sclerosis, in

which myelin is lost, resulting in unreliable signal conduction by CNS axons and a breakdown in neuronal communication.Thus, remyelination of axons after repair is likely to be impor-tant for enhancing functional recovery. Research indicates thattransplanted cells can help remyelinate axons. Olfactory en-sheathing cells (which resemble Schwann cells), oligodendro-cytes, and oligodendrocyte precursor cells all have been dem-onstrated to be effective in providing myelin for axons in vivoafter experimentally induced demyelination (9, 17). A recentstudy, in fact, reported that adult stem cells injected intrave-nously can remyelinate CNS axons (41), potentially providinga simple and convenient cell delivery method without requir-ing intracranial access. Cell transplantation currently is being

tested in clinical trials as a treatment for demyelinating dis-eases (52). These advances suggest that axon remyelinationafter MEMS-assisted repair also may be achieved using a cellreplacement strategy.

Assessment of Successful Repair

During surgery, an immediate assessment of whether therepair on a particular pair of axons is successful provides anongoing tabulation of the number of repairs accomplished.One convenient test of axonal function is the demonstration ofelectrical conduction from the host axon into the donor axonthrough the fusion site. Because electrode arrays are already in

place for axon manipulation and axon electrofusion, it should be possible to use different elements within the array close tothe axon of interest to act as stimulating and recording elec-trodes. This assessment of electrical conduction can be prepro-grammed to occur after each electrofusion, and the informa-tion will allow an objective intraoperative scoring of repairsuccess.

Mechanical Support after Axon Repair

A number of considerations suggest that an engineeredmicrostructure acting as a supporting scaffold may be usefulin the postrepair period. Recent studies have examined the

suitability of various polymers and biogels for use as scaffoldsin promoting axon regeneration (45). After axon repair, theseeding of cells for axon myelination will benefit from a three-dimensional structure providing a means of cell localizationand a substrate for cell anchoring. Furthermore, the polymericor biogel scaffolds can be infiltrated with chemicals and sub-stances that enhance axon viability and recovery, encouragevascularization, and decrease inflammation as well as scar-ring. Last, the engineered scaffold can be used to providemechanical support.





SUMMARY

In this article, we propose a new paradigm for the treatmentof nerve injuries based on the direct repair of damaged axons.This repair methodology uses a novel combination of micro-technology, electrokinetic axon manipulation, and the well-

established biological principle of cell fusion. These threefields of research have been integrated in a multidisciplinaryapproach to develop a solution for a significant clinical prob-lem that currently has no specific treatment. The findingsreported here provide some initial proof of principle for thecore technologies we intend to use for axon repair. This newtreatment paradigm for nerve injuries raises a number ofclinical issues such as defining the optimal time, number, andtype of axons for repair. Although existing knowledge sug-gests possible answers to some of these questions, solutions toother problems will have to come from additional research before microtechnology-based axon repair can become a pos-sibility in patients. It is hoped that each advance in axon repair

technology will spur additional research to provide us with acomprehensive understanding on how best to pursue neuro-surgical intervention at the microscale level.

REFERENCES

1. Aguayo AJ, Clarke DB, Jelsma TN, Kittlerova P, Friedman HC, Bray GM:

Effects of neurotrophins on the survival and regrowth of injured retinal

neurons. Ciba Found Symp 196:135–144; discussion 144–138, 1996.

2. Aguayo AJ, Rasminsky M, Bray GM, Carbonetto S, McKerracher L, Villegas-

Perez MP, Vidal-Sanz M, Carter DA: Degenerative and regenerative re-

sponses of injured neurons in the central nervous system of adult mammals.

Philos Trans R Soc Lond B Biol Sci 331:337–343, 1991.

3. Aldskogius H, Kozlova EN: Central neuron-glial and glial-glial interactionsfollowing axon injury. Prog Neurobiol 55:1–26, 1998.

4. Basso DM, Beattie MS, Bresnahan JC: Descending systems contributing to

locomotor recovery after mild or moderate spinal cord injury in rats: Ex-

perimental evidence and a review of literature. Restor Neurol Neurosci

20:189–218, 2002.

5. Bates G, Saunders J, Sowers A: Electrofusion: Principles and applications, in

Sowers A (ed): Cell Fusion. New York, Plenum Press, 1987, pp 376–395.

6. Berns MW, Tadir Y, Liang H, Tromberg B: Laser scissors and tweezers.

Methods Cell Biol 55:71–98, 1998.

7. Biering-Sorensen F: Urinary tract infection in individuals with spinal cord

lesion. Curr Opin Urol 12:45–49, 2002.

8. Bisby M: Regeneration of peripheral nervous system axons, in Waxman S,

Kocis J, Stys P (eds): The Axon: Structure, Function, and Pathophysiology. New

York, Oxford University Press, 1995, pp 553–578.

9. Blakemore WF, Franklin RJ: Transplantation options for therapeutic central

nervous system remyelination. Cell Transplant 9:289–294, 2000.

10. Bradbury EJ, Moon LD, Popat RJ, King VR, Bennett GS, Patel PN, Fawcett SB

McMahon JW: Chondroitinase ABC promotes functional recovery after

spinal cord injury. Nature 416:636–640, 2002.

11. Caroni P, Schwab ME: Antibody against myelin-associated inhibitor of

neurite growth neutralizes nonpermissive substrate properties of CNS white

matter. Neuron 1:85–96, 1988.

12. Chen DF, Schneider GE, Martinou JC, Tonegawa S: Bcl-2 promotes regen-

eration of severed axons in mammalian CNS. Nature 385:434–439, 1997.

13. Coumans JV, Lin TT, Dai HN, MacArthur L, McAtee M, Nash C, Bregman

BS: Axonal regeneration and functional recovery after complete spinal cord

transection in rats by delayed treatment with transplants and neurotrophins.

J Neurosci 21:9334–9344, 2001.

14. Detrait ER, Yoo S, Eddleman CS, Fukuda M, Bittner GD, Fishman HM:

Plasmalemmal repair of severed neurites of PC12 cells requires Ca(2) and

synaptotagmin. J Neurosci Res 62:566–573, 2000.

15. Eysel UT, Wolfhard U: The effects of partial retinal lesions on activity and

size of cells in the dorsal lateral geniculate nucleus. J Comp Neurol 229:

301–309, 1984.

16. Fouad K, Dietz V, Schwab ME: Improving axonal growth and functional

recovery after experimental spinal cord injury by neutralizing myelin asso-ciated inhibitors. Brain Res Brain Res Rev 36:204–212, 2001.

17. Franklin RJ: Remyelination by transplanted olfactory ensheathing cells.

Anat Rec 271B:71–76, 2003.

18. Fuhr G, Glasser H, Muller T, Schnelle T: Cell manipulation and cultivation

under a.c. electric field influence in highly conductive culture media.

Biochim Biophys Acta 1201:353–360, 1994.

19. Gascoyne P, Pethig R, Satayavivad J, Becker FF, Ruchirawat M:

Dielectrophoretic detection of changes in erythrocyte membranes following

malarial infection. Biochim Biophys Acta 1323:240–252, 1997.

20. Gascoyne PR, Vykoukal J: Particle separation by dielectrophoresis. Electro-

phoresis 23:1973–1983, 2002.

21. Godell CM, Smyers ME, Eddleman CS, Ballinger ML, Fishman HM, Bittner

GD: Calpain activity promotes the sealing of severed giant axons. Proc Natl

Acad Sci U S A 94:4751–4756, 1997.

22. Goldberg JL, Klassen MP, Hua Y, Barres BA: Amacrine-signaled loss of

intrinsic axon growth ability by retinal ganglion cells. Science 296:1860–1864, 2002.

23. Goslin K, Banker G: Experimental observations on the development of

polarity by hippocampal neurons in culture. J Cell Biol 108:1507–1516, 1989.

24. Guillery RW: Quantitative studies of transneuronal atrophy in the dorsal

lateral geniculate nucleus of cats and kittens. J Comp Neurol 149:423–438,

1973.

25. He Z, Koprivica V: The Nogo signaling pathway for regeneration block.

Annu Rev Neurosci 27:341–368, 2004.

26. Heida T, Rutten WL, Marani E: Dielectrophoretic trapping of dissociated

fetal cortical rat neurons. IEEE Trans Biomed Eng 48:921–930, 2001.

27. Heida T, Rutten WL, Marani E: Experimental investigation on neural cell

survival after dielectrophoretic trapping. Arch Physiol Biochem 110:373–

382, 2002.

28. Hui SW, Stenger DA: Electrofusion of cells: Hybridoma production by

electrofusion and polyethylene glycol. Methods Enzymol 220:212–227, 1993.

29. Kandel E, Schwartz J, Jessell T (eds): Principles of Neural Science. New York,McGraw-Hill, 2000.

30. Karsten U, Stolley P, Seidel B: Polyethylene glycol and electric field-

mediated cell fusion for formation of hybridomas. Methods Enzymol 220:

228–238, 1993.

31. Liu CY, Spicer M, Apuzzo MLJ: The genesis of neurosurgery and the

evolution of the neurosurgical operative environment: Part II—Concepts for

future development, 2003 and beyond. Neurosurgery 52:20–35, 2003.

32. Lodish H, Berk A, Matsudaira P, Kaiser CA, Krieger M, Scott MP, Zipursky

L, Darnell J: Molecular Cell Biology, 5th ed, W.H. Freeman, 2004.

33. Loy DN, Talbott JF, Onifer SM, Mills MD, Burke DA, Dennison JB, Fajardo

LC, Magnuson DS, Whittemore SR: Both dorsal and ventral spinal cord

pathways contribute to overground locomotion in the adult rat. Exp Neurol

177:575–580, 2002.

34. Ludwin S: Pathology of the myelin sheath, in Waxman S, Kocis J, Stys P

(eds): The Axon: Structure, Function, and Pathophysiology. New York, Oxford

University Press, 1995, pp 412–437.

35. Malessy MJ, Bakker D, Dekker AJ, Van Duk JG, Thomeer RT: Functional

magnetic resonance imaging and control over the biceps muscle after

intercostal-musculocutaneous nerve transfer. J Neurosurg 98:261–268, 2003.

36. Mano Y, Nakamuro T, Tamura R, Takayanagi T, Kawanishi K, Tamai S,

Mayer RF: Central motor reorganization after anastomosis of the musculo-

cutaneous and intercostal nerves following cervical root avulsion. Ann

Neurol 38:15–20, 1995.

37. Merkler D, Metz GA, Raineteau O, Dietz V, Schwab ME, Fouad K: Loco-

motor recovery in spinal cord-injured rats treated with an antibody neutral-

izing the myelin-associated neurite growth inhibitor Nogo-A. J Neurosci

21:3665–3673, 2001.

SRETAVAN ET AL.




38. Neil GA, Zimmermann U: Electrofusion, in Duzgunes N (ed): Methods in

Enzymology. New York, Academic Press, 1993, pp 174–196.

39. Okabe M, Ikawa M, Kominami K, Nakanishi T, Nishimune Y: ‘Green mice’

as a source of ubiquitous green cells. FEBS Lett 407:313–319, 1997.

40. Petersen KE: Silicon as a mechanical material. Proc IEEE 70:420–457, 1982.

41. Pluchino S, Quattrini A, Brambilla E, Gritti A, Salani G, Dina G, Galli R, Del

Carro U, Amadio S, Bergami A, Furlan R, Comi G, Vescovi AL, Martino G:

Injection of adult neurospheres induces recovery in a chronic model ofmultiple sclerosis. Nature 422:688–694, 2003.

42. Polla DL, Erdman AG, Robbins WP, Markus DT, Diaz-Diaz J, Rizq R, Nam

Y, Brickner HT, Wang A, Krulevitch P: Microdevices in medicine. Annu Rev

Biomed Eng 2:551–576, 2000.

43. Rafuse VF, Gordon T: Self-reinnervated cat medial gastrocnemius muscles.

I. Comparisons of the capacity for regenerating nerves to form enlarged

motor units after extensive peripheral nerve injuries. J Neurophysiol 75:

268–281, 1996.

44. Roy S, Ferrara LA, Fleischman AJ, Benzel EC: Microelectromechanical sys-

tems and neurosurgery: A new era in a new millennium. Neurosurgery

49:779–799, 2001.

45. Schmidt CE, Leach JB: Neural tissue engineering: strategies for repair and

regeneration. Annu Rev Biomed Eng 5:293–347, 2003.

46. Schucht P, Raineteau O, Schwab ME, Fouad K: Anatomical correlates of

locomotor recovery following dorsal and ventral lesions of the rat spinal

cord. Exp Neurol 176:143–153, 2002.47. Schwab ME: Repairing the injured spinal cord. Science 295:1029–1031, 2002.

48. Shigley J, Mischke C, Budynas R: Mechanical Engineering Design. New York,

McGraw-Hill, 2003.

49. Silver J, Miller JH: Regeneration beyond the glial scar. Nat Rev Neurosci

5:146–156, 2004.

50. Smith DH, Wolf JA, Meaney DF: A new strategy to produce sustained

growth of central nervous system axons: continuous mechanical tension.

Tissue Eng 7:131–139, 2001.

51. Somogyi J, Eysel U, Hamori J: A quantitative study of morphological reor-

ganization following chronic optic deafferentation in the adult cat dorsal

lateral geniculate nucleus. J Comp Neurol 255:341–350, 1987.

52. Stangel M, Hartung HP: Remyelinating strategies for the treatment of mul-

tiple sclerosis. Prog Neurobiol 68:361–376, 2002.

53. Stoppini L, Buchs PA, Muller D: Lesion-induced neurite sprouting and

synapse formation in hippocampal organotypic cultures. Neuroscience 57:

985–994, 1993.

54. Wakayama T, Perry AC, Zuccotti M, Johnson KR, Yanagimachi R: Full-term

development of mice from enucleated oocytes injected with cumulus cell

nuclei. Nature 394:369–374, 1998.

55. Wang X, Becker FF, Gascoyne PR: Membrane dielectric changes indicate induced

apoptosis in HL-60 cells more sensitively than surface phosphatidylserine expres-

sion or DNA fragmentation. Biochim Biophys Acta 1564:412–420, 2002.

56. Wang X, Yang J, Gascoyne PR: Role of peroxide in AC electrical field

exposure effects on friend murine erythroleukemia cells during

dielectrophoretic manipulations. Biochim Biophys Acta 1426:53–68, 1999.

57. Wilmut I, Schnieke AE, McWhir J, Kind AJ, Campbell KH: Viable offspring

derived from fetal and adult mammalian cells. Nature 385:810–813, 1997.

58. Yoshimura N, Smith CP, Chancellor MB, de Groat WC: Pharmacologic and

potential biologic interventions to restore bladder function after spinal cord

injury. Curr Opin Neurol 13:677–681, 2000.

59. You SW, Chen BY, Liu HL, Lang B, Xia JL, Jiao XY, Ju G: Spontaneous

recovery of locomotion induced by remaining fibers after spinal cord tran-

section in adult rats. Restor Neurol Neurosci 21:39–45, 2003.60. Zhai Q, Wang J, Kim A, Liu Q, Watts R, Hoopfer E, Mitchison T, Luo L, He

Z: Involvement of the ubiquitin-proteasome system in the early stages of

wallerian degeneration. Neuron 39:217–225, 2003.

61. Ziv NE, Spira ME: Induction of growth cone formation by transient and

localized increases of intracellular proteolytic activity. J Cell Biol 140:223–

232, 1998.

Acknowledgments

This study was supported by the That Man May See Foundationand the Sandler Family Supporting Foundation. No financial support

was received in conjunction with the generation of this manuscriptsubmission and none have any personal or institutional financialinterest in drugs, materials, or devices described in their submissions.Patent Pending to the Regents of the University of California formicrodevices and methods of axon repair.

COMMENTS

This is a well-written paper on an innovative new nerve repairparadigm that employs microscale neurosurgery to repair single

axons using micro/nanotechnology. The authors report a multidisci-plinary strategy to manipulate and operate at the single axon level anddiscuss the design of microfabricated devices for axon cutting, dielec-trophoresis (DEP), for axon manipulation, and electrofusion of axons.They also present interesting preliminary data to support their novelstrategy. As predicted by a brilliant article in Neurosurgery in 2003 thatforesaw that microtechnology and nanotechnology surely wouldmake important contributions to clinical neurosurgery in the future(1), this report is just one excellent example of research advances inthat direction. This type of creative thinking is exactly what is neededto tackle the difficult issues involved in the repair of neural injuries.There currently is no technique available for determining acutelywhich closed peripheral nervous system injuries will recover sponta-neously and which will require surgical intervention. We will needviable answers to many such basic questions before this type oftechnology can reach clinical testing.

In our lab, we have successfully elongated axons of both human andrat dorsal root ganglia in vitro with a computer-controlled mechanicaldevice and started transplantation experiments using in vitro grown GFPlabeled axons into host animals with acute sciatic nerve injuries (2, 3). Itis conceivable that the technology described by Dr. Sretavan et al. couldtake advantage of our in vitro elongated axons as donor axons in therepair of acutely injured axons. Obviously, there are numerous technicalhurdles that need to be overcome before this strategy can be applied to

patients, but we concur with the authors that this report will stimulateadditional research in this direction to provide neurosurgeons and neu-roscientists with a better understanding of how to advance neurosurgicalintervention at the microscale and nanoscale.

Jason H. Huang

Eric L. Zager

Philadelphia, Pennsylvania

1. Liu CY, Spicer M, Apuzzo ML: The genesis of neurosurgery and the evolu-

tion of the neurosurgical operative environment: Part II—concepts for future

development, 2003 and beyond. Neurosurgery 52:20–33, 2003.

2. Huang JH, Zager EL, Pfister BJ, Smith DH: Human Dorsal Root Ganglia

Neurons as Live Nerve Constructs: Implications for Transplantation. Pre-

sented at the AANS Annual Meeting, Orlando, Florida, May 2004.3. Huang JH, Groff RF, Zhang J, Pfister BJ, Zager EL, Smith DH: Living Nerve

Constructs Bridging Peripheral Nerve Lesions Demonstrate Long-term Sur-

vival and Restoration of Function. Platform Presentation (373.7) at the 34th

Annual Meeting of the Society for Neuroscience, San Diego, California,

October 2004.

Dr. Stretavan et al. have presented a wonderful glimpse into thefuture of “high tech” Neurosurgery. In this regard, they have ex-

ploited microtechnology to its fullest current potential. Although, whatthey have presented seems like “rocket science” bordering on “sciencefiction” to most, their observations, accomplishments, and technological





developments are truly insightful. At the outset, most would suggest thatwhat the authors ultimately propose, clinically meaningful and usefulsurgery on axons, is impossible or even ludicrous. However, the strategyand methodologies they outline should lead us to believe that some daythey (or those who they intrigue) will actually “pull it off.”

Dr. Stretavan et al. have introduced several unique strategies and have

suggested a global strategy for their implementation. These include 1) atechnique for axon cutting with a suspension mounted silicon nitrideknife; 2) a technique for axon manipulation, dielectrophoresis (DEP), thatcauses the movement of small polarizable objects, such as neurons in anonhomogeneous alternating current electrical field; 3) axon electrofu-sion techniques; 4) the confirmation of electrophysiologically successfulelectrofusion; 5) a surgical platform for accomplishing the aforemen-tioned; and 6) a mechanism to complete such tasks in “bulk” (parallelprocessing) so that hundreds, thousands, or millions of axons could berepaired in a reasonable amount of time.

What Dr. Stretavan et al. have proposed is not “science fiction.”They are sitting at the forefront of, and are hopefully going to guideus into, this most exciting of neurosurgical frontiers. For this, they areto be heartily congratulated.

Edward C. BenzelCleveland, Ohio

This report illustrates and discusses the concept of repairing damagedaxons by combining three separate microtechniques into a single

surgical platform. Sequentially, these include an axon micro-cutting de-vice made from silicon with a 20-nanometer tip, electrodes capable ofnoncontact manipulation of individual axons using DEP, and the fusionof aligned axon segments by delivering a pulse of electricity through thesame electrodes used for DEP. They provide illustrations depicting thecutting of axons, as well as the electro-fusion of overlapping, adjacentaxon segments. However, the precise (two-dimensional) alignment of cutaxons to be fused using DEP is not shown. Perhaps this is because it is notfeasible with the technology currently available.

Although the authors do not provide evidence of a successful axon

spliced in vitro using their proposed surgical platform, we believe itshould be readily possible. However, the important question remainsas to whether it can be applied to the central or peripheral nervoussystem in vivo. As with other technologies, if adequate resources areavailable and commercial interest is strong enough, a clinically appli-cable prototype may eventually appear. As the authors allude, axonrepair could be computer automated, with even multiple surgicalplatforms being simultaneously applied to an injury site. For effi-ciency, the process would utilize automated, real-time recognition ofthose axon terminals that are in the need of repair. This could beaccomplished with visual pattern recognition software.

One of the more obvious barriers to its eventual use would be thepreparation and alignment of axons in vivo prior to repair. Chemicalskeletonization may be one option, as the authors point out. But do theynecessarily need to be skeletonizedto such a degree? Considering that we(us and the computers) can probably identify cut axon terminals on end,even in their connective tissue environment, then one may extrapolatethat with proper alignment and electro-fusion, axon repair could occurwithout extensive, and possible damaging, skeletonization. Once pre-pared, axon alignment in three-dimensional space would likely be re-quired, not only in a two-dimensional plane as described in this report.

Even if axon repair becomes possible, the question remains whetherthe distal axon segment will remain viable after repair. To our knowl-edge, there have been no studies addressing this. Furthermore, whathappens to their terminal connections? It is plausible that with a quickenough repair these distal axon segments will not undergo degener-

ation. More information is required. An experiment using the larger,and thus more readily manipulated, squid axon may be a good

starting point. For example, the squid axon’s functional outcome can be investigated following transection and electro-fusion at various

times. We believe that the window for this type of axon repair may bequite short, and therefore, it require pharmacological adjuvants to

prolong this potential state in which axon salvage is possible.We commend the authors on their innovative and thought-

provoking application, which is a great example of how cross-disciplinary collaboration allows us to achieve new heights in the field

of neurosurgery.

Stephen M. Russell

Patrick J. Kelly

New York, New York

Dr. Sretavan et al. publish what can be considered the next era innerve repair: nanotechnology. The authors applied microscale

knives to resect an axon, and, with the aid of an electrical field, were

able to interpose a ‘healthy‘ piece of axon, or graft, between theproximal and distal stumps of a divided axon. This technique is

promising for peripheral nerve and spinal cord regeneration.Although the authors postulate multiple ways of achieving these

goals, there are many hurdles that still need to be overcome. They feelthat with these techniques, 180 axons could be repaired within an hour.

Performing this on an injured patient at a ‘nano-level‘ to a major periph-eral nerve with about 20,000 fibers would require many hours and also

would be difficult in deeply seated nerves or spinal cord in the depths ofthe spinal cord with surrounding spinal fluid. These considerations pro-

vide only challenges forfuture work and do not negate the importance ofthis contribution. Restoring electrical impulses in these axons is what is

going to make the concept work. Whether this can be accomplishedalong with remyeliniation is topic for future investigation.

Although this now may seem to be a far-fetched concept, the futuremay see a more practical application of this new technology, which

will translate into better outcomes for our patients. We wholeheart-edly congratulate the authors for their results and look forward to

further developments.

Rashid M. Janjua

David G. Kline

New Orleans, Lousiana

In this paper, the authors describe an intriguing approach to axonalrepair. They have taken advantage of advances in microtechnology to

consider the direct repair of damaged single axons by substituting seg-

ments of healthy donor axons. The basic steps to this mode of axon repairare described, along with prototypes of surgical devices that may bring

this concept to reality in the future. Without a doubt, the development ofneurosurgery has been marked by a progressive minimalism. The surgi-

cal paradigm described here may represent yet another step in thisdirection. At the present time, considerable development in technologies

mayeventually allow manipulation and “surgery” at even smaller scales.Although the concept of single axon repair may seem fantastic, future

realities enabled by the emerging field of nanotechnology may be evenmore astonishing. Irrespective of the eventual clinical utility of the de-

vices proposed in this work, I applaud the authors for their creativity andhope that work such as this serve to stimulate all neurosurgeons. I read

this paper with great enthusiasm.

Charles Y. Liu

Los Angeles, California

SRETAVAN ET AL.


Documents

David W. Sretavan et al- Microscale Surgery on Single Axons