Living myofibroblast–silicon composites for probingelectrical coupling in cardiac systemsMenahem Y. Rotenberga,1, Naomi Yamamotob, Erik N. Schaumannb, Laura Matinoc,d, Francesca Santoroc,and Bozhi Tiana,b,e,1

aThe James Franck Institute, The University of Chicago, Chicago, IL 60637; bDepartment of Chemistry, The University of Chicago, Chicago, IL 60637; cTissueElectronics, Center for Advanced Biomaterials for Healthcare, Istituto Italiano di Tecnologia, 80125 Naples, Italy; dDepartment of Chemical Materials andIndustrial Production Engineering, University of Naples Federico II, 80125 Naples, Italy; and eThe Institute for Biophysical Dynamics, The University ofChicago, Chicago, IL 60637

Edited by John A. Rogers, Northwestern University, Evanston, IL, and approved September 26, 2019 (received for review August 8, 2019)

Traditional bioelectronics, primarily comprised of nonliving syntheticmaterials, lack cellular behaviors such as adaptability and motility.This shortcoming results in mechanically invasive devices andnonnatural signal transduction across cells and tissues. More-over, resolving heterocellular electrical communication in vivo isextremely limited due to the invasiveness of traditional intercon-nected electrical probes. In this paper, we present a cell–siliconhybrid that integrates native cellular behavior (e.g., gap junctionformation and biosignal processing) with nongenetically enabledphotosensitivity. This hybrid configuration allows interconnect-freecellular modulation with subcellular spatial resolution for bioelec-tric studies. Specifically, we hybridize cardiac myofibroblasts withsilicon nanowires and use these engineered hybrids to synchronizethe electrical activity of cardiomyocytes, studying heterocellularbioelectric coupling in vitro. Thereafter, we inject the engineeredmyofibroblasts into heart tissues and show their ability to seam-lessly integrate into contractile tissues in vivo. Finally, we apply localphotostimulation with high cell specificity to tackle a long-standingdebate regarding the existence of myofibroblast–cardiomyocyteelectrical coupling in vivo.

optical stimulation | cardiomyocytes | myofibroblast | silicon nanowires |electrical coupling

Electronic and optoelectronic systems for in vivo cellular in-terrogation require minimally invasive tissue access within

the 3-dimensional tissue volume. Micropipette electrodes (1, 2),microelectrode arrays (3, 4), and field effect transistors (5–7)have enabled numerous single-cell studies, although they aretypically substrate-bound devices. Optogenetics can offer mech-anistic insights into cellular processes, but it requires geneticmodification (8–10), which limits its potential clinical applica-tions. Recent developments in flexible implantable devices haveyielded biointerfaces (11–15) that are minimally invasive and canintegrate wireless electronics and optoelectronics with opto-genetics. Currently, the material tool kit is primarily based onsynthetic components. When interfacing with live and dynami-cally changing tissues, seamless integration of the device is limitedby the remaining mechanical invasiveness of the materials andnonnatural biological signal transduction at the biointerfaces. Wepropose that living hybrid systems with dynamic and developingbehaviors can offer new opportunities for bioelectric interfacesdue to the adaptability and motility of the cellular componentsand the diverse physical properties of the materials components.Silicon nanowires (SiNWs) can be spontaneously internalized

by many cell types (16), which allows for hybridization whileavoiding abrasive electroporation, sonication, or genetic modifi-cation. Following internalization, cell–SiNW hybrids may be har-vested using standard cell culture techniques for both in vitro andin vivo downstream applications (Fig. 1A). The light scatteringfrom SiNWs distinguishes the hybrids from surrounding SiNW-free cells, eliminating the need for fluorescent labeling. Usinga focusing laser, the cell–SiNWs hybrids may be photostimulated

via photoelectric (17) and photothermal (18) mechanisms. As thephotostimulation depends on the colocalization of light and theSiNW, a localized stimulus can be applied deep into the tissue.Moreover, their freestanding nature minimizes their invasivenessdue to persistent in vivo mechanical mismatch.In this work, we apply our cell–SiNW hybrid system to the in-

vestigation of intracellular electrical coupling in cardiac systems.Hybridizing SiNWs with myofibroblasts (MFs), we demonstratethat this living hybrid tool can be used for investigating intercel-lular electrical coupling in vitro and in vivo. Using the MF–SiNWhybrid tool to compare MF–MF electrical coupling with MF–cardiomyocytes (CMs) coupling in vitro, we detect 2 differentcalcium flux propagation mechanisms—one for amplified CMpropagation and the other for passive MF propagation. We reportthat, unlike bare SiNWs, our MF–SiNW hybrids can be seamlesslyintegrated into contractile cardiac tissue. Finally, we use our livinghybrid tool to tackle the long-standing debate regarding the ex-istence of in vivo MF–CM heterocellular coupling.

In Vitro Electric Coupling and Optical PacingCoaxial SiNWs were synthesized as previously reported (17, 19).Scanning electron microscope (SEM) images of the SiNWs show a1-dimensional geometry with a ∼300-nm diameter and a core-shell

Significance

Techniques for intracellular electrical interrogation are chal-lenging to perform in vitro and are extremely limited in vivo,especially inside the tissue volume. Here, we hybridized cellswith silicon nanowires that transduced laser illumination intoelectrical interrogation in a simple, local, and leadless manner.We used these hybrids to address the question of whethercardiomyocytes and myofibroblasts electrically couple in vivo,which is important for understanding the contribution of scartissue to abnormal electrical conduction in the heart. Wefound that although myofibroblasts were able to couple tocardiomyocytes in vitro, no such coupling was observed in vivo.This suggests that the degree of heterocellular electrical couplingin the whole heart is far less substantial than in cell culture.

Author contributions: M.Y.R. and B.T. designed research; M.Y.R., N.Y., and E.N.S. per-formed research; L.M., F.S., and B.T. contributed new reagents/analytic tools; M.Y.R.,N.Y., E.N.S., and B.T. analyzed data; and M.Y.R. and B.T. wrote the paper.

The authors declare no competing interest.

This article is a PNAS Direct Submission.

Published under the PNAS license.

Data deposition: The raw data that was used to generate Figs. 1–4 have been depos-ited on figshare (https://figshare.com/articles/Data_from_Living_myofibroblast_silicon_composites_for_probing_electrical_coupling_in_cardiac_systems/9968294).1To whom correspondence may be addressed. Email: hrotenberg@uchicago.edu orbtian@uchicago.edu.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1913651116/-/DCSupplemental.

First published October 17, 2019.

www.pnas.org/cgi/doi/10.1073/pnas.1913651116 PNAS | November 5, 2019 | vol. 116 | no. 45 | 22531–22539

ENGINEE

RING

Dow

nloa

ded

by g

uest

on

May

29,

202

1

morphology, as visible in a randomly broken SiNW (Fig. 1B). Wethen hybridized the free-standing SiNWs with primary MFs viaspontaneous internalization (16) (Fig. 1C). We also verified MFs’viability by a live/dead assay, which showed negligible effect of theinternalized SiNW on MFs’ viability (SI Appendix, Fig. S1). Toevaluate whether our MF–SiNW composites can enable electricalcoupling in CMs in vitro, we optically stimulated a MF–SiNWhybrid cell in a coculture of CMs and MF–SiNWs and visualizedthe responses using the calcium-sensitive dye Fluo-4. In this set-ting, we applied a laser pulse focused at an SiNW within a MF–SiNW hybrid and monitored the effect of the photostimulation on

neighboring CMs. The internalized SiNWs display random distri-butions of lengths, diameters, and locations. Therefore, it is diffi-cult to determine a clear stimulation threshold. However, byapplying laser stimulations of different durations (1 ms and 5 ms)on the same SiNWs, we determined that longer pulses were morelikely to elicit an action potential (AP) in neighboring CMs (Fig.1D). The threshold value of ∼5 μJ per pulse is comparable to thatin the previously reported optical extracellular stimulation ofneurons, where a 5.4-μJ threshold was identified (17). However, itis worth mentioning that we did not obtain the 1-to-1 AP per lightpulse response, as was achieved for the neuromodulation (17). The

C

0.0

0.2

0.4

0.6

0.8

1.0

Stimulation success rate

1ms

5ms

SiNW-1 SiNW-2

Prob

abilit

y

0 50 100 150 200 250 300 350 400 4500

1

2

3

4

5Photo-stimulation

ROI 3

ROI 2

ROI 1

Time [s]In

terv

al b

etw

een

AP [s

]

F

A

Objective

Objective

SiNWs

Fibroblast

Cardiomyocyte

Bright field

2

3

Fluo-4

1

Before pacing While pacing

E

0

349 ms

B

D

n-1

n-2

n-1

n-2

0 5 10 15 20 25 30 35 40

dF/F

410 420 430 440 450

Pulse

ROI1

G

Fig. 1. In vitro electric coupling and optical pacing. (A) Illustration of the proposed MF–SiNW hybrid methodology. SiNWs are seeded on MFs and allowed tohybridize. The MF–SiNW hybrids can be harvested and cocultured with CMs or injected into heart tissue, where they provide high-resolution photomodulation.(B) SEM images of coaxial p-i-n SiNWs. (Top) One-dimensional morphology of an SiNW. (Bottom) Representative cross-section of a randomly broken SiNW, wherethe coaxial feature is clear. (Scale bars, 100 nm.) (C) Confocal images of cytoplasmatic (calcein AM, green) and membranal (CellMask, red) staining show that SiNWsare internalized by MFs. The SiNWs (white) are detected by reflected light. Yellow dashed lines represent 2 cross-sectioned z-n slices. (Scale bars, 10 μm.) (D) Longerpulse durations (1 ms or 5 ms, 1 mW) for SiNW stimulation increase the likelihood of provoking an AP in neighboring CMs. Results shown for 2 different SiNWs.Error bars represent SE of the mean from >20 stimulations for each SiNW. (E) Effect of photomodulation on MF–SiNW/CM coculture. (Top Left) Bright-field imageshows perinuclear arrangement of SiNWs within MF; arrowhead indicates stimulated SiNW. (Top Right) Fluorescent Ca2+ imaging in analyzed ROIs 1 to 3. (BottomLeft) Heat map of AP propagation before photopacing shows unsynchronized CM beating in ROI 1 and 3, and no electrical activity in ROI 2. (Bottom Right) Following∼400 s of photopacing (5 ms, 1 mW, 1.3 Hz), all regions are completely active and synchronized. Field of view: 169 μm × 169 μm. (F) Summary of activity in all ROIsfollowing optical stimulation of MF–SiNW hybrid. Results are plotted as the average of the time intervals between consecutive APs for each ROI. Black line, pacingrate of laser pulse. (G) dF/F vs. time of electrical activity of ROI 1; initial slow rate of electrical activity gradually increases and synchronizes with the laser pulses.

22532 | www.pnas.org/cgi/doi/10.1073/pnas.1913651116 Rotenberg et al.

Dow

nloa

ded

by g

uest

on

May

29,

202

1

intracellular location of the nanowires in the cells prevented usfrom further increasing the laser power without damaging the cell,while an extracellular configuration as used in the neuro-modulation study (17) is more tolerant of light-triggered damage.Although it has been previously reported that MFs can mediateelectrical activity between CMs in vitro (20–26), the ability of ourMF–SiNW hybrid system to manipulate natural bioelectric signalsin MFs and consequently modulate the electrical activity ofneighboring CMs has not been reported.We then asked if optical modulation of MF–SiNW hybrids

could induce overdrive pacing in cocultured CMs. The bright-fieldcontrast in Fig. 1 E, Top Left shows a perinuclear arrangement ofSiNWs within an MF–SiNW hybrid, with the stimulated SiNWmarked with a red arrow. Within this field, we selected 3 regions ofinterest (ROIs), each containing different CMs (Fig. 1 E, TopRight). Optical mapping of the calcium flux before stimulationshowed that ROIs 1 and 3 were spontaneously active, but thetiming of the contractions was not synchronized (Fig. 1 E, BottomLeft and Movie S1). As pacing requires repetitive stimulations, welowered the laser power to ∼1 mW and paced the MF–SiNWhybrid at 1.3 Hz. Upon optical pacing, we observed a gradual in-crease in the ROI contraction rates. Fig. 1F plots the time intervalsbetween consecutive contractions for each ROI; although imme-diate overdrive pacing was not observed, the contraction frequencygradually increased and approached the target pacing rate (1.3Hz). Moreover, the optical stimulation also induced synchroniza-tion of the contractions in the different ROIs (Fig. 1 E, BottomRight and Movie S1). As electrical coupling depends on the celldensity and the location of the CMs relative to the stimulated MF–SiNW hybrid, the observed response to the stimulation differedamong the ROIs. The contraction rate of ROIs 2 and 3, whichwere adjacent to the stimulated hybrid, immediately increased(Fig. 1F) upon stimulation. ROI 1, which was not in direct contactwith the target hybrid, gradually synchronized to the target pacingrate (Fig. 1 F andG) with a slower response rate due to mediationof the signal by other cells (see SI Appendix, Fig. S2 for full dF/Ftrace). We also performed pacing with higher laser power, whichresulted in an immediate, short-lived response, probably due todamage to the cells (SI Appendix, Fig. S3). Moreover, we showedthat CMs can be paced to different frequency (∼0.75 Hz; SI Ap-pendix, Fig. S4). These phenomena were observed in 5 indepen-dent cell cultures.

Investigation of In Vitro Heterocellular Electrical CouplingUsing the MF–SiNW HybridTo demonstrate the utility of our hybrid tool to investigate in vitrointercellular electrical coupling, we optically stimulated an MF–SiNW hybrid in coculture and compared the resulting effect inneighboring MFs and CMs. Fig. 2A shows a coculture of cell–SiNW hybrids and CMs loaded with calcium-sensitive dye. Base-line recording of the cells revealed that 3 cells were spontaneouslybeating CMs, while the others were static MFs. Initially, thespontaneous AP propagation had a specific directionality (Fig. 2 B,Left). Upon optical stimulation, however, a local calcium flux wasinitiated at the stimulation site and all 3 neighboring CMs wereimmediately activated, with an AP propagating radially from thestimulated hybrid (Fig. 2 B, Right). Thereafter, a calcium fluxslowly propagated through the MFs, as illustrated by mappingthe long-term effect of the same optical stimulation (Fig. 2C, SIAppendix, Fig. S5, and Movie S2). Interestingly, the slow calciumflux through the MF–SiNWs seems to propagate entirely in-dependently of the AP propagating through the CMs. This cell-specific response suggests that optical stimulation of the MF–SiNW activates 2 different mechanisms through which thecalcium flux propagates through the coculture. We then used apure MF culture, in which we stimulated different composites tobetter characterize the MF–MF coupling. Fig. 2D and Movie S3show 2 representative results from the stimulation of 2 different

MF–SiNW hybrids; the yellow arrows indicate the differentdirectionality and signal propagations originating from the stimu-lated SiNWs. These results demonstrate our ability to control theorigin of the stimulation with cell specificity and high spatial res-olution using our cell–SiNW hybrid.To quantify calcium propagation between and within different

cell types, we used a computer algorithm (Materials and Methodsand SI Appendix, Figs. S6–S8). Fig. 2E illustrates the propagationvelocities 1) from the stimulated MF–SiNW hybrid to neighboringMFs (MF–MF), 2) within each MF (MF–intracellular), and 3)from the stimulated MF–SiNW hybrid to neighboring CMs (MF–CM). The fastest calcium propagation velocity (average 988 μm/s)was from MF–SiNW hybrids to CMs. This was significantly fasterthan propagation between MFs (MF–MF) and within each MF(MF–intracellular) (P < 0.0001 for both). This large difference incalcium propagation velocity rates supports our hypothesis that 2different calcium flux propagation mechanisms exist in the co-culture—one for the amplified CM propagation and the otherfor passive MF propagation. Moreover, a closer look at MF in-tracellular velocities revealed a decay in the differential of the flowwith respect to the time of activation (Fig. 2F), meaning that MFsthat are further down the propagation path demonstrate slowercalcium flux propagation. We believe this calcium wave propagatesfreely through the cytosol to the borders of the MF; thereafter, itpropagates to the neighbor MF through gap junctions (20, 21) ortunneling nanotubes (27), a bottleneck that slows down thepropagation (Fig. 2G). This hypothesis explains 2 observed phe-nomena in our results on cultured MFs: The first is that the overallintercellular calcium wave propagation velocity, which combinesthe fast intracellular (MF–intracellular) and slow intercellular(MF–MF) propagation, is slower than the average intracellularMF calcium flux velocity (Fig. 2E), and the second is that the flowdecays with time after the stimulation (Fig. 2F). To investigatewhether the electrical coupling is mediated by gap junctions, weused carbenoxolone to completely block connexin 43. Indeed,upon stimulation of coculture with blocked gap junctions, onlythe stimulated cell was affected (SI Appendix, Fig. S9). We thenshowed that this effect was reversible, as changing the media tocarbenoxolone free media showed that intercellular coupling wasrestored (SI Appendix, Fig. S9). We also performed immunocyto-chemistry staining for connexin 43 (SI Appendix, Fig. S10). Thedegree of connexin 43 expression between adjacent CMs wasclearly higher than that between CMs and MFs, which was in turnhigher than that between adjacent MFs. Overall, these results fitwell with the accepted mechanism in which MFs passively mediateelectrical coupling (20, 21).

In Vivo Seamless Integration of the MF–SiNW HybridTo use our hybrids to perform in vivo electrical interrogation withcell specificity and high spatial resolution, we first demonstratethat they can form a seamless integration with the contractilecardiac tissue. To show this, we injected either MF–SiNW hybridsor bare SiNWs into the left ventricular (LV) wall of a transplantedheart (Movie S4) to assess integration into cardiac tissue (Fig. 3A).Histological analysis, performed 2 to 5 d postprocedure, revealedhybrid MFs adjacent to healthy and striated native CMs (Fig. 3Band SI Appendix, Fig. S11). In hearts which had received bareSiNWs, we observed that the interface was encapsulated by fibrotictissue that was significantly thicker than the tissue from the MF–SiNW-injected hearts (Fig. 3 B and C and SI Appendix, Fig. S11).In rare cases where bare SiNWs were near CMs, the CMs werevisibly weakened, lacking striation. These bare SiNW-induced ef-fects were accompanied by a dramatic increase in levels of theimmune response-related proteins CD3 and CD11b (Fig. 3E andSI Appendix, Fig. S12), likely indicating direct irritation by theSiNWs and an inflammatory response. Hematoxylin and eosinstaining (Fig. 3D and SI Appendix, Fig. S13) and Masson’s tri-chrome stain (SI Appendix, Fig. S14) further validated the

Rotenberg et al. PNAS | November 5, 2019 | vol. 116 | no. 45 | 22533

ENGINEE

RING

Dow

nloa

ded

by g

uest

on

May

29,

202

1

formation of thick fibrotic tissue by the bare SiNWs. Nuclearstaining with a membrane-impermeable dye (propidium iodide; SIAppendix, Fig. S15) showed no difference in the number of dead

cells between MF–SiNW hybrid hearts and bare SiNW hearts. Theabsence of dead cells in the bare SiNW hearts implies that anydead cells were likely removed and replaced by the fibrotic

Spontaneous AP

0

9.5 s

0

137 ms

CMCMMF

CM

MF

MF

MFs-CMs propagation

MFs-MFs propagation

A B

C D

E

1 2

FCalcium flow vs. excitation time

Tmax [s]0 2 4 6 8

0.0

0.2

0.4

0.6

0.8

1.0

δ(ν)

[ 1/s

]

CD1D2

0

8 s

GCalcium flow

**

MF-MF

MF-Intra

cellu

lar

MF-CM

020406080

100

500

1000

1500

2000

V elo

city

[μm

/s]

Laserpulse

Proposed model

Free diffusionFree diffusion

Bottleneck

Gap-junctionor Nanotube

HighCa2+

LowCa2+

Fast propagation

Slow propagation Slow propagation

Fig. 2. Investigation of in vitro heterocellular electrical coupling using the MF–SiNW hybrid. (A) Fluorescent image of MF–SiNW hybrid (white arrowhead)and neighboring MFs (blue dashed lines) and CMs (red dashed lines) analyzed in B and C. Field of view: 169 μm × 169 μm. (B, Left) Heat map showing a spon-taneous AP in CMs with specific directionality (yellow arrows). (B, Right) Heat map showing that optical stimulation of MF–SiNW hybrid results in a faster AP thatpropagates with different directionality to 3 adjacent CMs (yellow arrows). Field of view: 169 μm × 169 μm. (C) At a later time (9.5 s), a slow calcium flux (from thesame optical stimulation) propagates through the MFs, independent of the optically induced APs in the CMs. Field of view: 169 μm × 169 μm. (D) Stimulation of 2different MF–SiNW hybrids (red arrows) results in propagation of 2 different calcium waves. Field of view: 169 μm × 169 μm. (E) Calcium wave propagation fromMFs to CMs (MF–CM) is significantly faster than propagation through neighboring MFs (MF–MF) or within MFs (MF–intracellular) (*P < 0.0001). Boxes representquartiles, and whiskers represent minimum and maximum value (n = 42, 26, and 9 for MF–MF, MF–intracellular, and MF–CM, respectively). (F) MFs further downthe propagation path demonstrate a decay in the calcium flux. (C, D1, and D2 correspond to C and D). (G) Proposed model for passive propagation within andbetween cells. Free passive diffusion is slowed by the bottleneck of intercellular gap junctions or tunneling nanotubes.

22534 | www.pnas.org/cgi/doi/10.1073/pnas.1913651116 Rotenberg et al.

Dow

nloa

ded

by g

uest

on

May

29,

202

1

tissue over the 2 d following the transplant procedure. Theseobservations indicate that the MF–SiNW hybrids are capableof seamlessly integrating in situ with cardiac tissue more ef-fectively than bare SiNWs, supporting their use as a livingbioelectric probe for studying MF–CM coupling.

Investigation of In Vivo Heterocellular Electrical CouplingUsing the MF–SiNW HybridIn vitro electrical coupling between MFs and CMs is well estab-lished and has been extensively studied (20–26). However, theextent to which in vivo coupling between these 2 cell types occurs isstill under debate and has only been studied indirectly (21, 28–33).We applied our MF–SiNW system, with its capacity for subcellularresolution, to this question. After the MF–SiNW hybrid integratedinto the cardiac tissue, we harvested the heart to perform ex vivoinvestigation of the heterocellular coupling. Live tissue slices were

cut with a vibratome (Fig. 4A and Movie S5), loaded with calcium-sensitive dye, and photostimulated via confocal microscopy. In thissetting, the reflective nature of the SiNWs allowed us to easilydetect the hybrid cells using transmitted light imaging (Fig. 4 B,Top) without genetic modifications. Optical mapping followinglaser stimulation (Fig. 4 B, Bottom) showed that the calcium fluxpropagated a short distance (<25 μm), affecting the stimulated celland perhaps an adjacent cell. Although this does not entirely ruleout the presence of gap junctions between the hybrids and nativetissue, it does show that electrical coupling is far less substantialin vivo than in vitro. A contextual understanding of this result isimportant as theoretical models indicate that a substantial numberof coupled MFs with a high degree of connectivity are required toinduce significant arrhythmogenic effects (34, 35). Our findingsseem to contradict recent studies which provided evidence thatCM–MF coupling exists in vivo (29, 30). However, such coupling

0

20

40

60

80

100MF-SiNW hybrid

Bare SiNWs

5 days2 days

*

*

Sca

r thi

ckne

ss [u

m]

Donorheart

Abdominalincisions

Recipient rat

In vivo integrationof NW or

MF-SiNW hybrid

Tissuecryosectioning

and IHC

MF-SiNW hybrid Bare SiNWA B

C

NW orMF-SiNW

hybrid injection

PFA fixation

MF-SiNW hybrid Bare SiNW

CD

-3C

D-1

1b

D

E

MF-SiNW hybrid Bare SiNW

Striation Deteriorated tissue

Fig. 3. In vivo seamless integration of the MF–SiNW hybrid. (A) Schematic of in vivo MF–SiNW experiment. Following transplantation, the donor heart is injectedwith MF–SiNW hybrids (5 × 104) or bare SiNWs (normalized to be half of total SiNW amount within hybrids). After 2 to 5 d, hearts are removed and fixed forimmunohistochemistry (IHC). (B) IHC images of tissues injectedwithMF–SiNWhybrids or bare SiNWs. (Left) Hybrids are adjacent to native well-striated CMs (green);no fibrotic encapsulation is observed. (Right) Bare SiNWs (white) are encapsulated by fibrotic tissue and are not found near healthy CMs; closest CMs (green) lackstriation and appear deteriorated. (Scale bars, 10 μm.) (C) Encapsulating scar thickness in hearts with MF–SiNW hybrids is significantly lower than in hearts with bareSiNWs (*P < 0.001). Two-day time point: n = 9 injections, n = 3 hearts; 5-d time point: n = 6 injections, n = 2 hearts). Error bars represent SE of the mean from n > 20measurements. (D) Hematoxylin and eosin staining of the hybrids adjacent to well-striated CMs (Left) and bare SiNWs in the detreated tissue (Right). (Scale bars,50 μm.) (E) IHC images of immune markers CD-11b and CD-3 show the low immune response to the hybrids as opposed to the bare SiNWs. (Scale bars, 100 μm.)

Rotenberg et al. PNAS | November 5, 2019 | vol. 116 | no. 45 | 22535

ENGINEE

RING

Dow

nloa

ded

by g

uest

on

May

29,

202

1

was evident only at the scar–myocardial border with no propaga-tion into the scar (31). When we compare our observed in vivophotostimulation effect with our observed in vitro effect, and withreported in vitro effects from previous studies which showed thatMFs relay signals up to distances of 300 μm (20), it is evident thatdirect coupling between MFs and other cells (MFs or CMs) is farless substantial in vivo. Even if the stimulated hybrid was not ad-jacent to native CMs, one would expect coupled MFs to propagatethe effect to the interfaced tissue. To verify the tissue slice viability,we plotted the dF/F-vs.-time profile of nearby CMs. As a positivecontrol, we recorded the effect of a standard electrical stimulationon the CMs immediately before applying the optical stimulation(Fig. 4 C, Top); a clear calcium flux resulted from electricalstimulation, confirming that the tissue was viable and electricallyactive. Immediately afterward, we recorded the effect of the laserpulse (Fig. 4 C, Bottom), which showed no response to the stim-ulation. Thus, the lack of response to optical stimulation may beattributed to weak coupling between MF–SiNW hybrids and thetissue. To further validate this, we also performed immunohisto-chemistry staining for connexin 43. Although this alone cannotrule out the existence of connexin 43 at MF–MF and MF–CMinterfaces (28), the fact that the MF region (SI Appendix, Fig. S16)seems to lack connexin signal further supports our conclusion.Another important consideration is the cell integration durationthat is needed for gap-junction formation in vivo. Although we

have seen heterocellular electrical coupling in vitro occurringwithin 1 d of coculture, it is possible that in vivo coupling requireslonger time. We tested integration duration of 2, 5, and 9 d post-transplantation, but there was no calcium propagation for tissuesamples collected up to 9 d (SI Appendix, Fig. S17B).Another aspect that needs to be considered is the fact that the

native cardiac tissue functions as a syncytium of CMs (36, 37);the interconnected and large CMs result in extremely low inputresistance and high capacitance. Therefore, the current thresholdfor sufficient depolarization is much higher in vivo than in vitro.However, this concern was addressed by previous work in whichoptogenetics was used for cardiac stimulation (38). Althoughhigher levels of optical power density were used in vivo, they werecomparable to that used in vitro (∼2:1 ratio). Therefore, in ourin vivo study, we started with 4 mW of laser pulse. When no effectwas obtained with 4 mW, we systematically increased the laserpower up to 85 mW in vivo (vs. ∼4 to 7 mW in vitro, ∼20:1 ratio),which also did not yield any electrical response. To verify that theoptical signal is not lost due to the relatively low temporal reso-lution of optical mapping, we also recorded an electrocardiograph(ECG) of the optically stimulated and spontaneously contractingtissue (Fig. 4D). Unlike the spontaneous APs recorded in the ECG(Fig. 4 D, Top), high-power laser pulses only resulted in a smallartifact in the ECG (Fig. 4 D, Bottom and SI Appendix, Fig. S17A).This artifact was likely due to light diffraction from the tissue

ECG recording

B

C D

0

1.3sTissueblock

Agar

Vibratome

Agar

Tissuesections

ECG + ECG -

MF-SiNW hybrid

Heartsection

Calcium sensitivedye loading

Obective

AHeart after

in vivo integrationof NW or

MF-SiNW hybrid

Photostimulations

0.1mV

1 s

Photo stimulation

Spontaneous

0.1dF/F

1 s

Calcium imaging

Electricalstimulations

Fig. 4. Investigation of in vivo heterocellular electrical coupling using the MF–SiNW hybrid. (A) Schematic of the ex vivo MF–SiNW experiment. Following in vivointegration of the MF–SiNW hybrids, the removed hearts were sectioned into live tissue slices using vibrating microtome. Live heart slices are placed in a confocalmicroscope and SiNWs within hybrids are photostimulated. (B, Top) Transmitted light image of MF–SiNW hybrid and surrounding tissue showing location of laserstimulation. (B, Bottom) Optical mapping shows that the laser-induced calcium flux does not propagate to the tissue. (Scale bars, 20 μm.) (C) Calcium imaging ofthe heart slice. (Top) Tissue is healthy and responsive to standard electrical stimulation (positive control). (Bottom) Optical stimulation has no effect on the sur-rounding tissue. Colors represent separate photostimulations and corresponding electrical positive controls. (D) Upon laser illumination, no effect is recorded onECG of the heart slice.

22536 | www.pnas.org/cgi/doi/10.1073/pnas.1913651116 Rotenberg et al.

Dow

nloa

ded

by g

uest

on

May

29,

202

1

heating the electrodes. We also performed whole-heart experi-ments to establish the tissues’ viability over a longer period (SIAppendix, Fig. S18). Despite the clear photoresponse shown asinduced movement of SiNWs, no electrical effect was observed onthe ECG recording (SI Appendix, Fig. S19). Taken together, ourresults suggest that while MF–CM coupling is observed in in vitrocultures it is significantly weaker in vivo.

OutlookIn summary, we have shown that hybridizing inorganic nano-materials with cells allows for seamless integration with nativetissue. The free-standing nature of our living, interconnect-free,and minimally invasive composite, along with silicon’s ability toabsorb tissue-penetrating near-infrared light, may give rise to fu-ture clinical applications. Our method allows precise cell-specificbioelectric induction with high spatial resolution and enables theoptical identification of hybrids. It is simple, straightforward, andrelies on standard optical microscopy with no need for sophisti-cated device fabrication or patch-clamp systems. We have dem-onstrated its utility in cardiac systems, as the contractile natureof the cardiac tissue poses a major challenge for establishing aseamless bioelectric interface from a mechanical prospective.However, our hybrid system may also be applied to many otherbiological scenarios where cell-specific interrogation and sub-sequent tracking of intercellular signal flow are required, suchas the study of neuronal connectomics (39–41). One could alsoenvision that the living cell–SiNW hybrid may be induced into apluripotent stem cell, which can later be differentiated intomany forms for nongenetic optical modulation.

Materials and MethodsMethods.Nanowire synthesis. Coaxial p-i-n-SiNWs were synthesized using an Aunanocluster-catalyzed chemical vapor deposition (CVD) process. Au colloidalnanoparticles (100-nm diameter; Ted Pella) were deposited onto Si substrates(Nova ElectronicMaterials) for use as catalysts. During the SiNWgrowth, silane(SiH4) was used as the Si reactant, diboron (B2H6, 100 ppm in H2) as the p-typedopant, phosphine (PH3, 1,000 ppm in H2) as the n-type dopant, and hydro-gen (H2) as the carrier gas. For the p-type core SiNW growth, SiH4, B2H6, andH2 were delivered at flow rates of 2, 10, and 60 standard cubic centimetersper min (sccm), respectively. P-type core SiNW growth was carried out for30 min at 470 °C and 40 torr. For the intrinsic Si shell (i-shell) deposition, thetemperature was ramped up to 650 °C, during which time no gas flow wasallowed, and vacuum was applied. Then, SiH4 and H2 were delivered at 0.3and 60 sccm, respectively, at 15 torr. For the n-type outer shell, PH3 gas wasadded at a flow rate of 1.5 sccm, under the same conditions. The shell de-positions were performed at 750 °C at a pressure of 20 torr for 15 min pershell. Both intrinsic and n-type shells were grown for 20 min.Cell culture.All animal procedures were approved by The University of ChicagoInstitutional Animal Care and Use Committee (IACUC) and conducted incomplete compliance with the IACUC Animal Care and Use Protocol. Heartswere excised from postnatal day 0 to 5 neonatal rats into ice-cold Hanks’Balanced Salt Solution (HBSS) without Ca2+ or Mg2+. Primary cardiac fibro-blasts and CMs were isolated using the Pierce Primary Cardiomyocyte IsolationKit (Thermo Fisher Scientific), according to the manufacturer’s protocol. Afterisolation, the suspended cells were preplated for 1 to 2 h, allowing the fi-broblasts to adhere to the tissue culture plate. The enriched CM populationwas seeded on fibronectin (Sigma)-treated glass bottom dishes. The fibro-blasts were allowed to proliferate in culture media (DMEM high glucose +10% FBS, 1% Glutamax and 1% penicillin–streptomycin) until cells reached∼80% confluency. As fibroblasts spontaneously differentiate into MFs instandard culture (42), we considered these cells to be MFs. The preplated MFswere then used for hybridization with SiNWs and coculture with CMs. Forseeding SiNWs, 9 mm2 of Si substrate with CVD-grown SiNWs was sonicatedfor 10 min in culture media and then seeded on MFs in a ∼55-cm2 dish (0.16-mm2 chip per cm2 culture). After 12 h, the culture was vigorously rinsed 5to 8 times until no free-floating SiNWs were observed. Partially internalizedSiNWs were allowed to complete internalization for 2 h. MF–SiNW hybridswere harvested via trypsinization for 2 min and then rinsed and centrifugedfor 5 min at 200 × g (low g was used to avoid damage to the cells due tomechanical stress and strain by the SiNWs). Harvested MF–SiNW hybrid cellswere reseeded alone, cocultured with CMs, or injected into hearts.

Live/dead assay. Cells were treated with different concentrations of SiNWs for12 h. Then, SiNWs were rinsed away and cells were loaded with LIVE/DEADViability/Cytotoxicity Kit, for mammalian cells (Thermo Fisher Scientific),which consists of calcein AM (4 μM) and ethidium homodimer-1 (2 μM) for30 min. Cell were imaged immediately after rinsing the dye off.In vitro optical stimulation. Cells (MFs or MFs–CMs coculture) were treated withcalcium-sensitive dye (2 μM Fluo-4, AM, cell permeant; Thermo Fisher Scientific)for 30 min at 37 °C. Cells were rinsed and incubated for 30 min to allowcomplete deesterification. The treated cells were then analyzed using aMarianas Yokogawa-type spinning disk confocal for visualizing and stimulat-ing the cells. The Marianas Yokogawa confocal system allows setting a stim-ulation point for a designated time in-between recorded time frame. However,the time duration for switching the optical shutter from recording to stimu-lation varied. Consequently, the resulting pacing rate had an SD of ∼6%. Thus,Fig. 1F shows the moving average of the pacing rate throughout the experi-ment. For blocking connexin 43, we treated the cells with 500 μM carbenoxolone(Apexbio) in DMEM for 30 min. After stimulation, the media was replacedwith fresh carbenoxolone free media to reverse the blocking effect.Immunocytochemistry. Cells (MFs or MFs–CMs coculture) were fixed (4%paraformaldehyde; Sigma) and permeabilized (0.2% Triton X-100; Sigma).The cells were blocked (2% bovine serum albumin) to prevent nonspecificbinding and incubated with rabbit anti-cardiac troponin I antibody (Abcam;for CMs), chicken anti-Vimentin antibody (Abcam; for MFs), and mouse anti-Connexin 43 antibody (Abcam). Cells were then incubated with Alexa Fluor488, Alexa Fluor 647, and Alexa Fluor 568 secondary antibodies (Abcam).ProLong Gold Antifade Mountant with DAPI (Themo Fisher) was used tolabel the nuclei and the cells were imaged using the Leica SP5 TandemScanner Spectral 2-Photon Confocal.Optical mapping. Fluo-4 videos were analyzed using ImageJ (43) and an onlineavailable macro (44) for creating ΔF/F movies. The videos were made intobinary masks according to a threshold manually selected as activated/notactivated, and the resulting stack was used to generate a time color code forthe activation propagation.

We measured the intracellular speed of calcium signals by tracking thewave front in each cell using a thresholded kymograph along the axis ofpropagation (SI Appendix, Fig. S6). At each time point, the position of thesignal front is given by the boundary between the thresholded and non-thresholded regions. To calculate the average speed of the signal along thelength of the kymograph, we fit a line by eye to the boundary, such that theslope of the line equals the speed.

To quantify the rate of intercellular propagation, we analyzed the opticalflow of ΔF/F movies. The optical flow was estimated with the Lucas–Kanademethod as implemented in the MATLAB Image Processing Toolbox (TheMathWorks). To identify the time of calcium activation in cells, we consideredthat the mean optical flow, <ν>(t), of a region enclosed by the boundaries ofthe cell is initially very low, undergoes a rapid increase alongside the fluo-4signal upon activation, and then gradually decays in the absence of any furtherstimulation (SI Appendix, Fig. S7, Top). For ease of visualization, the <ν> signalis normalized to the initial time point for each cell. We can then characterizethe activation time of a cell as the point where the differential of themean optical flow signal reaches its peak (SI Appendix, Fig. S7, Bottom)We smoothed the mean optical flow signal using a Gaussian filter with σ =3Δt,where Δt is the time interval between frames in the movie, and found themaximum of the function ΔÆνæðtÞ= νðt + 3ΔtÞ− Æνæðt− 3ΔtÞ as the differential,an analogy to the derivative. We adopted this latter step in order to accountfor the variation different cells showed in the time to reach the maximummean flow (Fig. 2F and SI Appendix, Fig. S8). Finally, we calculated the speed ofthe calcium signal between cells i and j as vCa2+ = ðtmax,j − tmax,iÞ=rij, where rij isthe distance between the centroid of the 2 cells.In vivo hybrid integration. All animal procedures were approved by The Uni-versity of Chicago IACUCand conducted in complete compliancewith the IACUCAnimal Care andUse Protocol. For cardiac transplantation,weused 2 rats (donorand recipient) per surgery. Following anesthetic induction and stabilization, thedonor animal’s chest and the recipient animal’s abdominal areas were shavedand aseptically prepared. For donor surgery, the chest was opened and theheart exposed. After isolation and ligation of the superior/inferior vena cavaand pulmonary vein, the aorta and pulmonary artery were cut and the heartwas collected. The heart was preserved in cold saline awaiting transplantation.For recipient surgery, the animal’s abdominal cavity was opened, and the aortaand vena cava were isolated and exposed. Using 10-0 nylon sutures, the do-nor’s aorta was connected to the recipient’s aorta and the donor’s pulmonaryartery was connected to the recipient’s inferior vena cava. Upon the comple-tion of transplantation, the muscle and chest peritoneum inner layers weresutured with 5-0 monofilament sutures. Subcutaneous tissue and skin weresutured with 5-0 monofilament nonabsorbable sutures. Immediately after

Rotenberg et al. PNAS | November 5, 2019 | vol. 116 | no. 45 | 22537

ENGINEE

RING

Dow

nloa

ded

by g

uest

on

May

29,

202

1

blood flow to the transplanted heart was resumed, we injected the hybrids (5 ×104 cells) or bare SiNWs (normalized to be half of the total amount of SiNWswithin the MF–SiNW hybrids, discussed below) into 3 different locations in theLV wall. After the procedure, the rats were treated daily with cyclosporin A(15 mg/kg) and methylprednisolone (2 mg/kg) to prevent immune rejection.After the designated time (2 to 9 d), the heart was excised from the recipientabdomen and used for ex vivo Langendorff perfused heart model (n = 4 forbare NWs and n = 6 for MF–SiNW hybrids) or sectioned to live tissue slices(n = 4 for bare NWs and n = 5 for MF–SiNW hybrids). Immunohistochemistrywas performed on the unstimulated parts of the injection site.

To measure the amount of SiNWs that were internalized by the MFs, wedrop-casted 50 μL of SiNWs suspension before seeding on the cells. There-after, to avoid coffee ring formations while drying, we mounted a coverslipand imaged the suspension immediately using dark-field microscopy (SIAppendix, Fig. S20). Then, after the SiNWs were internalized, we collected asample of the media with noninternalized SiNWs and repeated the pro-cedure. We then calculated the number of pixels that were positive forSiNWs (all frames were given the same threshold), which correlated to theamount of NWs in the suspension. The difference in SiNW-occupied areabetween the samples before and after introduction to cells trends roughlyproportional to the amount of internalized SiNWs. From this procedure, wefound that approximately half of SiNWs were internalized; as such, we usedhalf of the amount of NWs that were internalized as the control of bareSiNWs injected in vivo.Immunohistochemistry. Hearts were excised from the recipient rat’s abdomen.The LV wall was removed and embedded in optimal cutting temperaturecompound (Fisher Scientific). Samples were snap-frozen in isopentane usingdry ice and stored at –80 °C. The frozen tissue was cut into 10-μm slices usinga cryotome (Cryostat NX50). The tissue sections were processed for standardimmunohistochemistry (same antibodies as for in vitro cells, plus antibodiesagainst immune response proteins CD3 and CD11b). Other slices were pro-cessed for standard hematoxylin and eosin or Masson’s trichrome staining.Vibratome sectioning of live tissue slices. Following removal from the recipientrat’s abdomen, the heart was placed in ice-cold HBSS buffer, and the aortawas cannulated in preparation for use in a Langendorff setup. OxygenatedHepes-buffered Tyrode’s solution (containing 126 mM NaCl, 5.4 mM KCl,10 mM glucose, 10 mM Hepes, 1 mM MgCl2, 2 mM CaCl2, 1.2 mM MgSO4,and 0.39 mM NaH2PO4; bubbled with 99.5% O2; pH 7.3) was perfusedthrough the cannulated aorta. The perfusion was passed through a heatingcoil and bubble trap (Radnoti), and the heart was placed in a water-jacketedbeaker (Fisher Scientific) to maintain the temperature at 37 °C. Rhod-2 cal-cium dye (5 μM; Abcam), and Pluronic F-127 (Thermo Fisher Scientific) wereadded to the perfusion buffer and loaded into the hearts (45). After 20 minof dye loading, the heart was perfused with dye-free media for 30 min andmoved to ice-cold oxygenated buffer. The heart was cut into live tissueslices according to Watson et al. (46). Briefly, the LV was removed andglued (Medbond Tissue Glue; Stoelting Co.) to an agar substrate so that the

muscle fibers would be aligned to the blade. A sapphire blade was used ona Leica VT1000 S vibratome to slice the heart while immersed in ice-coldoxygenated buffer, and the slices were kept in the same cutting buffer forup to 3 h.Ex vivo optical stimulation of tissue slices. Slices of the heart were moved to aprewarmedmicroscope stage and allowed to equilibrate to 37 °C for 20 to 30minwhile bubbled with oxygen. A Leica SP5, STED-CW Superresolution LaserScanning Confocal was used to visualize the MF–SiNW hybrids within thetissue, and the STED setting was used to apply high-power optical stimu-lation. The stimulated slices were visualized to record calcium flux changes as aresult of optical stimulation. An SI-200 Stimulus Isolator (IWorx) was used toapply electrical stimulation as a positive control to verify slice viability. Toperform the ECG recording of optical stimulation and spontaneous con-tractions (not of electrical stimulation), we used an isolated Biopotentialrecording preamplifier (C-ISO-256; IWorx) and an amplifier (IA-400D; IWorx)that were connected to a digitizer (DI-1100; DataQ).Statistical analysis.Weused GraphPad Prism 8 to perform all statistical analysis.For propagation velocities we used 1-way ANOVA with Tukey’s multiplecomparisons test. For scar size we used unpaired t test. In all datasets, P <0.05 was considered significant.Data availability. The raw data that was used to generate Figs. 1 B–G, 2 A–F,3 B–E, and 4 B–D have been deposited on figshare (47). These data in-clude: raw SEM images (Fig. 1B); z-stack of confocal imaging (Fig. 1C); rawvideos of the photostimulations (sld files, Fig. 1D); raw video of the photo-stimulation, bright filed image of the SiNWs, fluorescent image of thefluo-4, and a text file with the stimulation timings (Fig. 1 E–G); raw videoof the photo-stimulation (Fig. 2 A–C); raw videos of the photostimulations(D1, D2, i, ii, iii, iv, v, and vi) (Fig. 2 D–F); z-stack of confocal imaging of thescar (Fig. 3B); images of the scar with the scar thickness as used for statis-tical analysis (Fig. 3C); Hematoxylin and Eosin images of the scar (Fig. 3D);z-stack of confocal imaging of the scar immune response (Fig. 3E); rawvideo of the photostimulation and the transmitted light image showingthe SiNWs (Fig. 4B); raw videos of the 3 different photostimulations andtheir intensity profiles (Fig. 4C); and ECG recording of the optical andelectrical stimulation (Fig. 4D).

ACKNOWLEDGMENTS. We thank Karen Watters for scientific editing of themanuscript; Prof. Yoram Etzion from Ben-Gurion University of the Negevand Prof. Ilya A. Fleidervish from Ben-Gurion University of the Negev fortheir scientific consultation and brainstorming; Prof. Margaret Gardel fromThe University of Chicago for her technical assistance; and the Human TissueResource Center at The University of Chicago, the Integrated Light microscopyCore Facility at The University of Chicago, and the Animal MicrosurgeryCenter (CORE) at The University of Chicago for their technical assistance. Thiswork is supported by the Air Force Office of Scientific Research (AFOSRFA9550-18-1-0503), US Army Research Office (W911NF-18-1-0042), US Officeof Naval Research (N000141612530, N000141612958), and the NationalInstitutes of Health (NIH NS101488)

1. B. Sakmann, E. Neher, Patch clamp techniques for studying ionic channels in excitablemembranes. Annu. Rev. Physiol. 46, 455–472 (1984).

2. K. Jayant et al., Targeted intracellular voltage recordings from dendritic spines usingquantum-dot-coated nanopipettes. Nat. Nanotechnol. 12, 335–342 (2017).

3. C. Xie, Z. Lin, L. Hanson, Y. Cui, B. Cui, Intracellular recording of action potentials bynanopillar electroporation. Nat. Nanotechnol. 7, 185–190 (2012).

4. J. T. Robinson et al., Vertical nanowire electrode arrays as a scalable platform forintracellular interfacing to neuronal circuits. Nat. Nanotechnol. 7, 180–184 (2012).

5. X. Duan et al., Intracellular recordings of action potentials by an extracellular nano-scale field-effect transistor. Nat. Nanotechnol. 7, 174–179 (2011).

6. B. Tian et al., Three-dimensional, flexible nanoscale field-effect transistors as localizedbioprobes. Science 329, 830–834 (2010).

7. Q. Qing et al., Free-standing kinked nanowire transistor probes for targeted intracellularrecording in three dimensions. Nat. Nanotechnol. 9, 142–147 (2014).

8. A. M. Packer, B. Roska, M. Häusser, Targeting neurons and photons for optogenetics.Nat. Neurosci. 16, 805–815 (2013).

9. E. S. Boyden, F. Zhang, E. Bamberg, G. Nagel, K. Deisseroth, Millisecond-timescale,genetically targeted optical control of neural activity. Nat. Neurosci. 8, 1263–1268(2005).

10. A. M. Packer, L. E. Russell, H. W. Dalgleish, M. Häusser, Simultaneous all-opticalmanipulation and recording of neural circuit activity with cellular resolutionin vivo. Nat. Methods 12, 140–146 (2015).

11. B. Tian, C. M. Lieber, Synthetic nanoelectronic probes for biological cells and tissues.Annu. Rev. Anal. Chem. (Palo Alto, Calif.) 6, 31–51 (2013).

12. D. H. Kim, R. Ghaffari, N. Lu, J. A. Rogers, Flexible and stretchable electronics forbiointegrated devices. Annu. Rev. Biomed. Eng. 14, 113–128 (2012).

13. K. J. Yu, Z. Yan, M. Han, J. A. Rogers, Inorganic semiconducting materials for flexibleand stretchable electronics. NPJ Flexible Electronics 1, 4 (2017).

14. S. P. Lee et al., Highly flexible, wearable, and disposable cardiac biosensors for remoteand ambulatory monitoring. NPJ Digit Med 1, 2 (2018).

15. G. Hong, C. M. Lieber, Novel electrode technologies for neural recordings. Nat. Rev.Neurosci. 20, 330–345 (2019).

16. J. F. Zimmerman et al., Cellular uptake and dynamics of unlabeled freestanding siliconnanowires. Sci. Adv. 2, e1601039 (2016).

17. R. Parameswaran et al., Photoelectrochemical modulation of neuronal activity withfree-standing coaxial silicon nanowires. Nat. Nanotechnol. 13, 260–266 (2018).

18. Y. Jiang et al., Rational design of silicon structures for optically controlled multiscalebiointerfaces. Nat. Biomed. Eng. 2, 508–521 (2018).

19. B. Tian et al., Coaxial silicon nanowires as solar cells and nanoelectronic powersources. Nature 449, 885–889 (2007).

20. G. Gaudesius, M. Miragoli, S. P. Thomas, S. Rohr, Coupling of cardiac electrical activityover extended distances by fibroblasts of cardiac origin. Circ. Res. 93, 421–428 (2003).

21. A. Klesen et al., Cardiac fibroblasts : Active players in (atrial) electrophysiology?Herzschrittmacherther. Elektrophysiol. 29, 62–69 (2018).

22. K. Goshima, Synchronized beating of and electrotonic transmission between myo-cardial cells mediated by heterotypic strain cells in monolayer culture. Exp. Cell Res.58, 420–426 (1969).

23. A. Hyde et al., Homo-and heterocellular junctions in cell cultures: An electrophysio-logical and morphological study. Prog. Brain Res. 31, 283–311.

24. K. Goshima, Formation of nexuses and electrotonic transmission between myocardialand FL cells in monolayer culture. Exp. Cell Res. 63, 124–130 (1970).

25. P. Kohl, A. G. Kamkin, I. S. Kiseleva, D. Noble, Mechanosensitive fibroblasts in thesino-atrial node region of rat heart: Interaction with cardiomyocytes and possiblerole. Exp. Physiol. 79, 943–956 (1994).

26. P. Camelliti, C. R. Green, P. Kohl, “Structural and functional coupling of cardiacmyocytes and fibroblasts” in Cardiovascular Gap Junctions, S. Dhein, Ed. (KargerPublishers, 2006), vol. 42, pp. 132–149.

27. K. He et al., Long-distance intercellular connectivity between cardiomyocytes andcardiofibroblasts mediated by membrane nanotubes. Cardiovasc. Res. 92, 39–47(2011).

22538 | www.pnas.org/cgi/doi/10.1073/pnas.1913651116 Rotenberg et al.

Dow

nloa

ded

by g

uest

on

May

29,

202

1

28. P. Kohl, R. G. Gourdie, Fibroblast-myocyte electrotonic coupling: Does it occur innative cardiac tissue? J. Mol. Cell. Cardiol. 70, 37–46 (2014).

29. T. A. Quinn et al., Electrotonic coupling of excitable and nonexcitable cells in theheart revealed by optogenetics. Proc. Natl. Acad. Sci. U.S.A. 113, 14852–14857 (2016).

30. M. Rubart et al., Electrical coupling between ventricular myocytes and myofibroblastsin the infarcted mouse heart. Cardiovasc. Res. 114, 389–400 (2018).

31. S. Nattel, Electrical coupling between cardiomyocytes and fibroblasts: Experi-mental testing of a challenging and important concept. Cardiovasc. Res. 114, 349–352 (2018).

32. A. M. De Mazière, A. C. van Ginneken, R. Wilders, H. J. Jongsma, L. N. Bouman, Spatialand functional relationship between myocytes and fibroblasts in the rabbit sinoatrialnode. J. Mol. Cell. Cardiol. 24, 567–578 (1992).

33. P. J. Lafontant et al., Cardiac myocyte diversity and a fibroblast network in thejunctional region of the zebrafish heart revealed by transmission and serial block-facescanning electron microscopy. PLoS One 8, e72388 (2013).

34. S. Rohr, Arrhythmogenic implications of fibroblast-myocyte interactions. Circ ArrhythmElectrophysiol 5, 442–452 (2012).

35. K. A. MacCannell et al., A mathematical model of electrotonic interactions betweenventricular myocytes and fibroblasts. Biophys. J. 92, 4121–4132 (2007).

36. S. Weidmann, The diffusion of radiopotassium across intercalated disks of mamma-lian cardiac muscle. J. Physiol. 187, 323–342 (1966).

37. W. C. De Mello, Effect of intracellular injection of calcium and strontium on cellcommunication in heart. J. Physiol. 250, 231–245 (1975).

38. T. Bruegmann et al., Optogenetic control of heart muscle in vitro and in vivo. Nat.

Methods 7, 897–900 (2010).39. A. Fornito, A. Zalesky, M. Breakspear, The connectomics of brain disorders. Nat. Rev.

Neurosci. 16, 159–172 (2015).40. H. Zeng, J. R. Sanes, Neuronal cell-type classification: Challenges, opportunities and

the path forward. Nat. Rev. Neurosci. 18, 530–546 (2017).41. J. W. Lichtman, J. Livet, J. R. Sanes, A technicolour approach to the connectome. Nat.

Rev. Neurosci. 9, 417–422 (2008).42. S. Rohr, Cardiac fibroblasts in cell culture systems: Myofibroblasts all along? J.

Cardiovasc. Pharmacol. 57, 389–399 (2011).43. C. T. Rueden et al., ImageJ2: ImageJ for the next generation of scientific image data.

BMC Bioinformatics 18, 529 (2017).44. J. Ackman, dFoFmovie-CatFullAutoSave.java. https://gist.github.com/ackman678/11155761.

Accessed 9 October 2019.45. K. Wang et al., Cardiac tissue slices: Preparation, handling, and successful optical

mapping. Am. J. Physiol. Heart Circ. Physiol. 308, H1112–H1125 (2015).46. S. A. Watson et al., Preparation of viable adult ventricular myocardial slices from large

and small mammals. Nat. Protoc. 12, 2623–2639 (2017).47. M. Y. Rotenberg, Data from: Living myofibroblast–silicon composites for probing

electrical coupling in cardiac systems. figshare. https://figshare.com/articles/Data_

from_Living_myofibroblast_silicon_composites_for_probing_electrical_coupling_in_cardiac_

systems/9968294. Deposited 10 October 2019.

Rotenberg et al. PNAS | November 5, 2019 | vol. 116 | no. 45 | 22539

ENGINEE

RING

Dow

nloa

ded

by g

uest

on

May

29,

202

1