Evidence of intercellular coupling between co-cultured adult rabbit ventricular myocytes and myofibroblasts


Corresponding author G. L. Smith: Institute of Biomedical and Life Sciences, West Medical Building, University of Glasgow, Glasgow G12 8QQ, UK. Email: g.smith@bio.gla.ac.uk


Intercellular coupling between ventricular myocytes and myofibroblasts was studied by co-culturing adult rabbit ventricular myocytes with previously prepared layers of cardiac myofibroblasts. Intercellular coupling was examined by: (i) tracking the movement of the fluorescent dye calcein; (ii) immunostaining for connexin 43 (Cx43); and (iii) measurement of intracellular [Ca2+] ([Ca2+]i). The effects of stimulating ventricular myocytes on the underlying myofibroblasts was examined by confocal measurements of [Ca2+]i using fluo-3. When ventricular myocytes were preloaded with calcein and co-cultured with myofibroblasts for 24 h, calcein fluorescence was detected in 52 ± 4% (n= 8 co-cultures) of surrounding myofibroblasts. Treatment with the gap junction uncoupler heptanol significantly reduced the movement of calcein (12 ± 3%, n= 6 co-cultures). Immunostaining showed expression of Cx43 in co-cultured myofibroblasts and myocytes. Field stimulation of ventricular myocytes co-cultured with myofibroblasts increased myofibroblast [Ca2+]i, no response was observed after treatment with heptanol or stimulation of fibroblasts in the absence of ventricular myocytes. Action potential parameters of ventricular myocytes in co-culture were similar to control values. However, application of the hormone sphingosine-1-phosphate (S-1-P) to the co-culture caused a depolarization of ventricular myocytes to approximately −20 mV. Sphingosine-1-phosphate had no effect on ventricular myocytes alone. Voltage-clamp measurements of isolated myofibroblasts indicated that S-1-P activated a significant quasi-linear current with a reversal potential of approximately −40 mV. In conclusion, this study shows that stimulation of the ventricular myocyte influences the intracellular Ca2+ of the linked myofibroblast via connexons. These intercellular links also allow the myofibroblasts to influence the electrical activity of the myocyte. This work indicates the nature of the gap junction-mediated bi-directional interactions that occur between ventricular myocyte and myofibroblast.

In addition to maintaining the extracellular matrix (ECM), cardiac fibroblasts secrete a number of growth factors and cytokines that may modulate the activity of adjacent fibroblasts and cardiac myocytes (Baudino et al. 2006). Electrical linkage between fibroblasts and myocytes from the atrium has been observed (Kohl et al. 2005), thereby implicating fibroblasts in the modulation of cardiac electrophysiology. In the sino-atrial node, fibroblasts express the gap junction protein connexin 40 (Cx40) in fibroblast-rich areas and the connexin 45 (Cx45) isoform where they intermix with atrial myocytes (Camelliti et al. 2004b). In the ventricle, the number of fibroblasts exceeds the number of ventricular myocytes (Nag, 1980), but the role of fibroblasts in the modulation of ventricular physiology is poorly understood.

Following a myocardial infarction (MI), a modified form of fibroblast termed ‘myofibroblast’ is common in the MI scar. Myofibroblasts differentiate from endogenous fibroblasts, infiltrating pericytes or stem cells (Baudino et al. 2006). They differ from fibroblasts in having greater expression of contractile proteins, in particular smooth muscle α-actin, and by the ability to secrete different factors (Powell et al. 1999). Within the MI scar, remaining ventricular myocytes coexist with myofibroblasts (Sun et al. 2002; Camelliti et al. 2004a; Li et al. 2005) and may be linked via gap junction proteins connexin 43 (Cx43) and Cx45 (Camelliti et al. 2004a). In vitro, following passaging, fibroblasts differentiate into myofibroblasts (Wang et al. 2005). Using this preparation, previous work has shown that adult ventricular myofibroblasts express both inwardly rectifying K+ currents and time- and voltage-gated K+ currents (Chilton et al. 2005; Shibukawa et al. 2005). In co-cultures of neonatal ventricular myocytes and fibroblasts, evidence of electrotonic coupling exists (Rook et al. 1992; Gaudesius et al. 2003; Miragoli et al. 2006). Furthermore, Cx43 and Cx45 staining was observed among myofibroblasts and between myofibroblasts and myocytes (Miragoli et al. 2006). However, electrotonic coupling has not been reported for adult ventricular myocytes and myofibroblasts. In the present study, adult rabbit ventricular myocytes were co-cultured for 24 h on a layer of adult rabbit ventricular myofibroblasts. Three independent approaches were used to demonstrate myocyte-myofibroblast coupling: (i) intercellular dye movement; (ii) the influence of stimulation of ventricular myocytes on myofibroblast intracellular [Ca2+] ([Ca2+]i); and (iii) the effect of activation of a ligand-gated ionic conductance in myofibroblasts on the electrical activity of ventricular myocytes. In this third aspect of the study, the effects of the hormone sphingosine-1-phosphate (S-1-P) were examined. Sphingosine-1-phosphate is released from platelets and macrophages in response to cell damage and is thought to be an important mediator of wound healing and angiogenesis (Pyne & Pyne, 2000). Levels of S-1-P in the nanomolar range are known to influence myofibroblast physiology (Pyne & Pyne, 2000) but have no direct effects on the electrophysiology of cardiac myocytes (MacDonell et al. 1998). This therefore provides an opportunity to investigate the influence of an agonist acting at the level of the myofibroblast on the activity of coupled cardiac myocytes.


Ventricular myocyte and myofibroblast isolation and culture

Cell isolation procedure complied with British Home Office regulations, and was approved by the University of Glasgow Animal Resources Centre. Rabbits were killed by an intravenous injection of 500 IU heparin together with an overdose of sodium pentobarbitone (100 mg kg−1). Isolated rabbit hearts were perfused retrogradely with 0.6 mg ml−1 collagenase (type 1, Worthington Chemical Co.) and 0.1 mg ml−1 protease (type XIV, Sigma Chemical Co.). Dissociated cells were centrifuged (10 min, 4 g), resuspended in Dulbecco's modified Eagle's Medium/F12 (DMEM) containing 10% fetal calf serum (FCS), penicillin (10 μ ml−1)/streptomycin (10 μg ml−1) and gentamycin (50 mg ml−1), and maintained under culture conditions (37°C, 5% CO2). Washing with DMEM/F12 the following day removed myocytes from the culture of fibroblasts. Myofibroblasts were used following first passage; freshly dissociated ventricular myocytes were added following 24 h serum starvation.

Dye coupling studies

Isolated ventricular myocytes were incubated with calcein AM (5 μmol l−1; 37°C, 30 min), washed, added to cultures of myofibroblasts, and left for 24 h. Heptanol (2 mmol l−1) was used in some co-cultures to block gap junction communication (Deleze & Herve, 1983; Burt & Spray, 1988; Rudisuli & Weingart, 1989; Niggli et al. 1989; Spray & Burt, 1990; Nelson & Makielski, 1991; Kimura et al. 1995). Calcein fluorescence was measured with a Bio-Rad Radiance 2000 confocal system. The total number of myofibroblasts surrounding a single myocyte within a 215 μm × 215 μm area was counted, and the portion of myofibroblasts with calcein fluorescence, indicative of dye-coupling, was expressed as a percentage of the total. Alternatively, myofibroblasts were loaded with calcein AM (37°C, 30 min) prior to addition of ventricular myocytes, and the percentage of ventricular myocytes with calcein fluorescence after 24 h co-culture was determined.

Immunostaining for Cx43

Myofibroblasts were seeded onto culture slides (Falcon). Freshly dissociated ventricular myocytes were added once myofibroblasts had achieved ≥ 60% confluence. Following 24 h in co-culture, cells were fixed (2% formaldehyde in NaCl–inorganic phosphate (Pi) solution, 30 min), permeabilized (1% Triton X-100 in NaCl–Pi solution, 10 min) and blocked (1% bovine serum albumin (BSA) in NaCl–Pi solution, 2 h). The cells were incubated overnight at 4°C with mouse anti-Cx43 monoclonal antibody (1:250 dilution). Cells were covered in foil and incubated for 1 h at room temperature with donkey fluorescein isothiocyanate (FITC)-conjugated antimouse IgG polyclonal antibodies (1:250 dilution). Fluorescence was monitored with an Olympus FV-1000 laser-scanning confocal microscope system.

Measurements of [Ca2+]i

Ventricular myocytes and myofibroblasts were incubated with fluo-3 AM (50 μmol l−1, 10 min) and superfused with Tyrode solution. Fluo-3 was excited using the 488 nm line of an Argon laser on a Bio-Rad Radiance 2000 laser-scanning confocal system. Emitted fluorescence above 515 nm was monitored using epifluorescence optics of a Nikon Eclipse inverted microscope with a Fluor ×60 water objective lens (numerical aperture 1.2). Optimal iris diameter provided an axial (Z) resolution of ∼0.9 μm, and X–Y resolution of ∼0.5 μm, based on full-width half-maximal amplitude measurements of images of 0.1 μm fluorescent beads (Molecular Probes). Data were acquired at 2 ms per line; pixel dimension was 0.42 μm (512 pixels per scan, 512 lines per frame). Single ventricular myocytes were field stimulated at 2 Hz and the fluo-3 fluorescence in the myofibroblasts within the surrounding 215 μm × 215 μm area was monitored at five frames per second.


Ventricular myocytes and myofibroblasts in co-culture, or monoculture, were superfused at 36–37°C in a MatTek Petri dish mounted on an inverted microscope (Diaphot 200, Nikon, UK). Myocyte action potentials recorded using the whole-cell patch-clamp technique were elicited by injecting 2–5 ms 1.2 × threshold current pulses (Axoclamp 2B amplifier and Digidata 1322A, Axon Instruments, union City, CA, USA) at 1 Hz. Myofibroblast currents were measured using the whole-cell patch-clamp method. Each cell was clamped at −80 mV and then stepped to −140 mV, after which it was depolarized in 10 mV, 1 s increments to 50 mV. Currents were recorded under control conditions, in the presence of S-1-P or vehicle, and following washout. In other experiments, myofibroblasts in monoculture were clamped at −90 mV and depolarized to 30 mV for 300 ms at a frequency of 1 Hz. This protocol was meant to approximate the changes in membrane potential associated with ventricular myocyte action potentials. Vehicle or S-1-P was applied (6 μl stock solution pipetted into the 3 ml bath; final concentrations from 0.2 fmol l−1 to 20 nmol l−1) and the effect of the vehicle or S-1-P on myofibroblast currents recorded. In a second set of these experiments, myofibroblasts in monoculture were incubated with fura-2 AM (5 μmol l−1) for 15 min at 37°C prior to initiation of the protocol. Fura-2 fluorescence signals were recorded at 100 Hz using a dual-wavelength spectrophotometric method previously described (Eisner et al. 1989).


Calcium-free Krebs–Henseleit buffer was used for cell isolation and contained (in mmol l−1): 130 NaCl, 5.4 KCl, 0.4 NaH2PO4, 3.5 MgCl, 5 Hepes, 20 taurine, 10 creatine and 11.1 glucose (pH 7.25 adjusted with NaOH, equilibrated with 100% O2). Tyrode superfusate contained (in mmol l−1): 144 NaCl, 5.4 KCl, 0.3 NaH2PO4, 1 MgCl2, 5 Hepes, 11.1 glucose and 1 CaCl2 (pH adjusted to 7.4 with NaOH). The patch-pipette solution contained (in mmol l−1): 12 NaCl, 20 KCl, 110 potassium aspartate, 1 CaCl2, 1 MgCl2, 4 K2ATP, 10 EGTA and 10 Hepes (pH adjusted to 7.2 with KOH). The NaCl–Pi solution contained (in mmol l−1): 140 NaCl, 2.7 KCl, 10 Na2HPO4 and 1.8 KH2PO4. All chemicals were obtained from Sigma except collagenase type 1 (Worthington Chemical Co.), BSA, FCS, gentamycin, penicillin, streptomycin (Gibco Invitrogen), fluo-3, calcein AM, fura-2 AM (Molecular Probes), S-1-P (Caymen Scientific), mouse anti-Cx43 monoclonal antibodies and donkey FITC-conjugated antimouse IgG polyclonal antibodies (Chemicon International).


Data are expressed as means ±s.e.m. Statistical significance was assessed by Student's unpaired t tests, in which P values of 0.05 were considered significant.


Myocyte–myofibrobast coupling: evidence from dye movement and connexin expression

Intercellular dye movement, indicative of functional intercellular coupling, was observed after pre-incubating ventricular myocytes with calcein AM and then, after washing off calcein AM, adding these myocytes to cultures of unloaded myofibroblasts. Calcein fluorescence spread from ventricular myocytes to the surrounding myofibroblasts within the 24 h co-culture period (Fig. 1). Figure 1Aa shows a transmission image of a ventricular myocyte on top of a field of myofibroblasts. In Fig. 1Ab and c, the calcein fluorescence image shows staining in both the ventricular myocyte and the surrounding myofibroblasts. On average, 52 ± 4% (n= 58 areas, 8 co-cultures) of myofibroblasts in a 215 μm × 215 μm area surrounding the preloaded myocyte showed calcein fluorescence (Fig. 1B, black bar) 24 h after addition of the myocyte. Co-incubation with heptanol (2 mmol l−1), which disrupts gap junction formation (Kimura et al. 1995), reduced the spread of calcein fluorescence to 12 ± 3% (n= 32 areas, 6 co-cultures) of the myofibroblasts (Fig. 1B, grey bar). No detectable transfer of calcein was recorded at the shorter incubation time of 4 h following addition of pre-incubated ventricular myocytes to the myofibroblast culture (0%, n= 30 areas, 2 co-cultures, Fig. 1B). The total number of myofibroblasts assessed in each case was 2677 (24 h), 1440 (24 h + heptanol) and 1676 cells (4 h). Myofibroblast–myocyte transfer of calcein was also obtained when myofibroblasts were pre-incubated with calcein AM and then unlabelled ventricular myocytes were placed in co-culture for 24 h (data not shown). Furthermore, immunostaining of myofibroblasts and myocytes following 24 h in co-culture revealed robust expression of Cx43 by both myofibroblasts and myocytes (Fig. 2). These results indicate that functional intercellular junctions have formed between ventricular myocytes and myofibroblasts following 24 h in co-culture.

Figure 1.

Evidence of calcein dye-coupling between adult rabbit myofibroblasts and ventricular myocytes in co-culture
Aa, representative transmitted light image of a myocyte pre-incubated with calcein AM following 24 h co-culture on a layer of myofibroblasts. Ab and c, calcein fluorescent images of an optical section through the middle of the myocyte (Ab) and an optical section from 10 μm below this (Ac), which contains only myofibroblasts. The position of the overlying ventricular myocyte is indicated with a grey outline. Scale bars represent 20 μm. B, mean ±s.e.m. (n) percentage of myofibroblasts exhibiting calcein fluorescence following 24 h co-culture under control conditions (black bar), with 2 mmol l−1 heptanol present in the culture media (grey bar), and within 4 h after addition of ventricular myocytes (right).

Figure 2.

C × 43 staining in co-cultured myocytes and fibroblasts
A, immunostaining for Cx43 was robust in myofibroblasts and ventricular myocytes maintained in co-culture for 24 h. Aa, immunofluorescent image showing FITC fluorescence, with Cx43 staining shown as punctate green spots. Ab, light micrograph of co-culture. Arrows indicate individual myocytes lying on a confluent field of myofibroblasts. Scale bar represnts 15 μm. Appropriate controls showed no non-specific staining. The areas indicated by the white dashed lines are shown expanded in Ba and b. The outer border of the ventricular myocyte is highlighted in both panels.

The influence of myocyte activity on myofibroblast [Ca2+]i

Given that ventricular myocytes and myofibroblasts exhibit intercellular coupling following 24 h in co-culture, it was of interest to determine whether these two cell populations were also functionally coupled. Accordingly, [Ca2+]i was monitored in 24 h co-cultured myofibroblasts and ventricular myocytes using the dye fluo-3 (Loughrey et al. 2003). Figure 3A shows a transmission image of a myocyte and its associated myofibroblasts (Fig. 3Aa), as well as a composite fluorescent image (Fig. 3Ab). During field stimulation of the ventricular myocyte, the intracellular [Ca2+] transiently increased in the myocyte in response to each stimulus. In a subset of the adjacent myofibroblasts, [Ca2+]i increased with a slow and sustained time course (Fig. 3B). As shown in Fig. 3B, [Ca2+]i increased in myofibroblasts 1 and 2 (Fig. 3B, light and medium grey) during the elevation of [Ca2+]i in the stimulated myocyte, while myofibroblast 3 appeared unaffected (Fig. 3B, dark grey). Figure 3C presents the average rise in [Ca2+]i observed in a group of 40 myofibroblasts during field stimulation of a single associated ventricular myocyte. Recovery of [Ca2+]i was observed in some myofibroblasts during the experimental period, with intracellular [Ca2+] declining at a rate of a rate of 0.135 ± 0.03 F/Fo units min−1 (where F/Fo is the ratio of fluorescence intensities; n= 9). The recovery of [Ca2+]i was slow compared with that observed in myofibroblasts after transient exposure to 1 mmol l−1 ATP (Fig. 3D). The peak intracellular Ca2+ signal (ΔF/Fo values) in field-stimulated ventricular myocytes increased by 0.79 ± 0.24 units (n= 6 myocytes) under co-culture conditions. However, the low frame rate available and the rapid kinetics of the ventricular myocyte mean that peak fluorescence values could not be obtained accurately. The transient nature of the Ca2+ signal meant that rise in mean fluorescence in myocytes during the stimulation period was small (ΔF/Fo= 0.19 ± 0.11, n= 6 myocytes). In contrast, the intracellular [Ca2+] in associated co-cultured myofibroblasts increased considerably more (ΔF/Fo= 0.61 ± 0.05, n= 91 cells, 3 co-cultures) in response to field stimulation. Direct comparisons are difficult because the basal fluorescence observed in the two cell types could be the result of different intracellular [Ca2+] levels. Co-incubation with heptanol (2 mmol l−1), which disrupts gap junction formation, inhibited the rise in Ca2+ within fibroblasts (ΔF/Fo=−0.03 ± 0.02, n= 62 cells, 5 co-cultures) during stimulation of the ventricular myocyte. Furthermore, when myofibroblasts in monoculture were subjected to field stimulation, intracellular [Ca2+] did not change significantly (ΔF/Fo=− 0.08 ± 0.02, n= 99 cells, 7 co-cultures), supporting the contention that field stimulation itself did not significantly change [Ca2+]i in myofibroblasts. Similarly, subthreshold field stimulation of ventricular myocytes in 24 h co-cultures was not associated with a rise in fluo-3 fluorescence in either the myocyte or the associated myofibroblasts (data not shown).

Figure 3.

Variation in fluo-3 fluorescence in myofibroblasts co-cultured with adult rabbit ventricular ventricular myocytes for 24 h
Aa, light micrograph showing a single myocyte after 24 h co-culture with myofibroblasts. A composite image of fluo-3 fluorescence is shown in Ab, in which ‘M’ refers to the myocyte. The three myofibroblasts which were recorded from are identified. B, fluo-3 intensity in the myocyte (black) and in individual myofibroblasts (grey lines) within the 215 μm2 area studied. C, average change in signal in 40 myofibroblasts associated with a single myocyte within a different 215 μm2 area. D, response of myofibroblasts to exogenous ATP (1 mm). The time periods over which the myocytes were field stimulated are denoted in B, C and D.

In principle, the observed increase of intracellular [Ca2+] in myofibroblasts during field stimulation of associated ventricular myocytes could have been produced by humoral factors released from the ventricular myocytes. To test this hypothesis, 1 mmol l−1 ATP was applied to monocultures of myofibroblasts. As shown in Fig. 3D, the response had a distinctly different time course from that seen in Fig. 3C. Addition of ATP caused a rapid increase of intracellular [Ca2+] followed by a slower decline, consistent with the characteristic changes in [Ca2+]i following activation of a Gq protein-coupled receptor (Brilla et al. 1998; Meszaros et al. 2000). The average initial increase in [Ca2+] following application of ATP (ΔF/Fo= 0.47 ± 0.06, n= 9 monocultures) was of comparable amplitude but quite different time course to that seen on myocyte stimulation. In the sustained presence of ATP, the intracellular Ca2+ decreased back to baseline and no further response to ATP could be elicited (data not shown), suggesting rapid receptor desensitization.

Effect of co-culture on myocyte electrophysiology

The possibility that co-cultured myofibroblasts and ventricular myocytes are electrically coupled was investigated by measuring passive subthreshold electrical responses, as well as action potential waveforms, in ventricular myocytes following 24 h in co- or monoculture. Ventricular action potentials were very similar in ventricular myocytes in co-culture and monoculture (Table 1), suggesting that intercellular coupling to surrounding myofibroblasts did not significantly affect action potential characteristics. Action potential durations at 50 and 90% repolarization (APD50 and APD90, respectively) were similar in co-cultured versus monocultured ventricular myocytes, as were diastolic potential (Vdiastolic) and peak depolarization (Vpeak; Table 1). Similarly, passive electrical properties were not statistically different between ventricular myocytes maintained in co-culture versus monoculture (Table 1). These data suggest that any electrophysiological effects on the ventricular myocyte of co-culturing with myofibroblasts are small compared with the error associated with single microelectrode measurements.

Table 1.  Electrical properties of ventricular myocytes following 24 h co-culture versus monoculture
 Co-cultured myocytesMonocultured myocytes
  1. Top 5 rows: mean ±s.e.m. (n) electrophysiological parameters of action potentials evoked by 1 Hz pacing in ventricular myocytes following 24 h co-culture or monoculture. Bottom 3 rows: mean ±s.e.m. (n) passive electrophysiological parameters evoked by injecting depolarizing or hyperpolarizing current in ventricular myocytes under co-culture or monoculture conditions. Abbreviations: APD50 and APD90, action potential durations at 50 and 90% repolarization, respectively; Vdiastolic, diastolic potential; and Vpeak, peak depolarization. ΔV is VdiastolicVpeak. Slope resistance (R) is based on the current response to a 10 mv hyper polarising pulse (−90 mv to −100 mv)

Vdiastolic (mV)−90 ± 1 (17)  −89 ± 1 (23)  
ΔV (mV)112 ± 6 (17) 100 ± 6 (23) 
Vpeak (mV)27 ± 1 (17)26 ± 1 (23)
APD50 (ms)252 ± 18 (17)263 ± 22 (23)
APD90 (ms)298 ± 19 (17)304 ± 25 (23)
Slope R (M±)12 ± 1 (17)13 ± 2 (23)
τon (ms) 1.8 ± 0.2 (17) 1.8 ± 0.2 (23)
τoff (ms) 3.0 ± 0.3 (17) 2.9 ± 0.3 (23)

Effect of S-1-P-induced myofibroblast current on myocyte electrophysiology

To investigate the influence of altered myofibroblast physiology on ventricular myocytes, the hormone S-1-P was applied to the co-culture system at concentrations of 0.2 fmol l−1 to 20 nmol l−1. Sphingosine-1-phosphate is released from macrophages and platelets during inflammation and is known to mediate changes in myobfibroblast physiology (Pyne & Pyne, 2000). When applied to monocultures of ventricular myocytes, S-1-P did not affect ventricular myocyte electrophysiology at concentrations up to 2 μmol l−1 (data not shown). Application of S-1-P (20 nmol l−1) to a co-culture caused rapid depolarization of ventricular myocyte membrane potential and loss of excitability (Fig. 4A and B). This effect could be partly reversed (Fig. 4A, right trace). Sphingosine-1-phosphate had no effect in monocultures of ventricular myocytes, in that membrane potential was −83 ± 0.6 mV (n= 7) under control conditions versus−82 ± 0.4 mV (n= 7) with S-1-P present; no effect on action potential duration was observed. On average, application of S-1-P affected 55 ± 8% (n= 13 co-cultures) of ventricular myocytes under co-culture with myofibroblasts, causing reversible depolarization from −83 ± 1 to −24 ± 2 mV (n= 20 cells). In separate experiments, pre-incubation of the myocyte–myofibroblast co-culture with heptanol (2 mmol l−1) for 5 min prior to addition of S-1-P blocked the effect on ventricular myocyte electrophysiology. In eight co-cultures with heptanol, S-1-P (40 nmol l−1) was ineffective at altering ventricular myocyte excitability (mean resting membrane potential in S-1-P + heptanol =−79 ± 0.8 mV, n= 8).

Figure 4.

Effects of 20 nmol l−1 sphingosine-1-phosphate (S-1-P) on action potential characteristics
Application of 20 nmol l−1 S-1-P during 1 Hz pacing depolarized and blocked excitability in a representative myocyte co-cultured with myofibroblasts (A, left trace, and B). The S-1-P-induced depolarization and loss of excitability were reversible following 10 min washout (A, right trace).

Effect of S-1-P on myofibroblast electrophysiology

In the absence of myocytes, the average membrane potential of cardiac myofibroblasts was −41 ± 6 mV (n= 11). On addition of 20 nmol l−1 S-1-P, the average membrane potential was not significantly different from control values (−33 ± 6 mV, n= 11). In voltage-clamped cardiac myofibroblasts (average capacitance 21 ± 1 pF, n= 30), the background conductance showed weak inward rectification (Fig. 5A). Addition of 20 nmol l−1 S-1-P activated a conductance with a relatively linear current–voltage relationship and a reversal potential of −38 ± 6 mV (n= 13, Fig. 5). Application of vehicle did not affect myofibroblast currents (data not shown). Single applications of 20 nmol l−1 S-1-P increased the inward current measured at −90 mV (approximate myocyte resting membrane potential) from −11 ± 6 to −75 ± 12 pA pF−1 and increased outward current at 30 mV (approximate peak of the action potential) from 24 ± 0.01 to 72 ± 11 pA pF−1 (n= 13, Fig. 5B). When myofibroblasts were clamped at −90 mV and stepped to 30 mV at 1 Hz, sequential application of S-1-P (from 0.2 fmol l−1 to 20 nmol l−1) increased the current in a concentration-dependent manner (Fig. 6). The concentration required for half-maximal activation of the S-1-P-induced current was 6.3 ± 3.6 pmol l−1 at 30 mV and 3.0 ± 1.4 pmol l−1 at −90 mV. Application of S-1-P (20 nmol l−1) was not associated with a change in fura-2 ratio, suggesting that S-1-P does not affect [Ca2+]i (Fig. 7). When myofibroblasts were clamped at −90 mV and stepped to 30 mV at 1 Hz (Fig. 7A), the fura-2 ratio signal was unchanged (Fig. 7C), both within each current pulse (Fig. 7B) and over the course of the protocol. Similarly, application of 20 nmol l−1 S-1-P during this voltage protocol did not influence fura-2 ratio (Fig. 7C), although a S-1-P-sensitive current was recorded (Fig. 7). The fura-2 ratio was 0.36 ± 0.01 (n= 25) during stimulation of the myofibroblasts with the voltage step and 0.37 ± 0.02 (n= 12) with S-1-P (20 nmol l−1) present.

Figure 5.

Sphingosine-1-phosphate activated a transmembrane ionic current in myofibroblasts
Application of 20 nmol l−1 S-1-P evoked an increase in the whole-cell currents in voltage-clamped myofibroblasts. A, representative current–voltage relationship under control conditions (^) and after 1–2 min 20 nmol l−1 S-1-P application (•). The inset panel is the difference current. Application of vehicle did not cause any change in myofibroblast current. B, transmembrane ion currents under control conditions (open bars) and evoked by 20 nmol l−1 S-1-P (black bars) at −90 (left) and 30 mV (right, means ±s.e.m., n= 13).

Figure 6.

Concentration dependence of the S-1-P-induced current change in myofibroblasts
A, representative traces from a monocultured myofibroblast voltage-clamped at −90 mV and then stepped repetitively to 30 mV for 300 ms at 1 Hz. Current changes recorded under control conditions (black line) and in response to increasing concentrations of S-1-P (grey lines) are shown. The response of a representative myofibroblast to S-1-P over the S-1-P concentration range (^= 30 mV; •=−140 mV) is shown in B, while mean increases in current are shown in C (means ±s.e.m., n= 11).

Figure 7.

Application of S-1-P and depolarization did not affect myofibroblast [Ca2+]i
A, myofibroblast membrane potential (in mV). Myofibroblasts in monoculture were depolarized at a rate of 1 Hz from −90 to 30 mV for 300 ms, mimicking the parameters of the average ventricular myocyte action potential. The time between sections is approximately 90 s. B, representative trace of myofibroblast current evoked by the step depolarization depicted in A, and with S-1-P application (20 nmol l−1; right trace). C, representative trace of myofibroblast fura-2 ratio prior to depolarization (left section), during ‘mock action potentials’ (middle section), and with 20 nmol l−1 S-1-P present (right section). Neither depolarization nor application of S-1-P affected myofibroblast fura-2 ratio.

In an attempt to reconcile the gradual increase in current caused by S-1-P application with the abrupt change in myocyte membrane potential observed in co-cultures, ventricular myocyte membrane potential was recorded under current-clamp conditions while stimulating action potentials at 1 Hz (Fig. 8). As shown in a representative trace in Fig. 8A, depolarizing current (top panel) was injected in increasing amounts while recording membrane potential (bottom panel). A gradual increase in inward current in the monocultured myocytes caused an abrupt change in diastolic potential, similar to that observed during application of S-1-P to co-cultured ventricular myocytes. As indicated in fig. 8, injection of 0.17 ± 0.02 nA (n= 22) of current was associated with sustained diastolic depolarization to −11 ± 3 mV (n= 22) and loss of excitability.

Figure 8.

Injection of a rectangular depolarizing current waveform depolarized and blocked excitability of ventricular myocytes in monoculture
In 24 h monocultures of ventricular myocytes, action potentials were evoked at 1 Hz by injection of 1.2 × threshold 10 ms current steps (A, top trace). A long depolarizing injection of current was then applied until diastolic membrane potential failed to repolarize and excitability was lost (A, bottom trace). As shown in B, the range of injected currents over which ventricular myocyte membrane potential remained depolarized was very narrow. Both panels show representative data from single ventricular myocytes. On average, injection of 0.17 ± 0.02 nA (n= 22) of current was associated with a sustained diastolic depolarization from −72 ± 1 to −11 ± 3 mV (n= 22). At this membrane potential, a loss of excitability in the monocultured myocyte was observed.


This is the first study to show that functional intercellular couplings between adult rabbit ventricular myocytes and cardiac myofibroblasts affect intracellular [Ca2+]i and cellular electrophysiology. The data indicate that intercellular coupling causes the [Ca2+]i within the myofibroblasts to be influenced by stimulation of the nearby ventricular myocyte. Furthermore, intercellular coupling allows the hormone S-1-P to alter ventricular myocyte electrophysiology indirectly by altering myofibroblast electrophysiology.

Myocyte–myofibroblast coupling: evidence from dye movement and connexin expression

In this study, detectable dye transfer between preloaded adult rabbit ventricular myocytes and myofibroblasts was not evident within 4 h of adding ventricular myocytes. Following 24 h in co-culture, 52 ± 4% of myofibroblasts were dye coupled with closely apposed myocytes (Fig. 1). In contrast, Driesen and co-workers reported dye-coupling as early as 1.5 h after adult rabbit cardiac fibroblasts were added to cultured adult rabbit ventricular myocytes (Driesen et al. 2005). The disparity of time course may arise from the different protocols used. In the present study, freshly dissociated adult ventricular myocytes were added to 60–95% confluent cultures of first passage myofibroblasts; 24 h later intercellular connections were evident. In the previous study, adult ventricular myocytes were maintained in culture for 24 h before adding freshly passaged fibroblasts.

The nature of the intercellular coupling observed in the present study between co-cultured adult ventricular myocytes and myofibroblasts is not known. However, it is likely that gap junctions formed between myocytes and myofibroblasts, and among myofibroblasts. Immunostaining detected expression of Cx43 in both myofibroblasts and myocytes following 24 h co-culture (Fig. 2). Punctate Cx43-positive features were observed over the surface of the myocyte; this is quite unlike the ribbon-like structures observed at the cell ends of adult myocytes. This pattern is typical for neonatal cardiomyocytes and suggests that the rabbit myocytes de-differentiate in culture while the links to the fibroblasts are established. Similar patterns of Cx43 staining were also reported in a previous study of co-cultured myocytes and fibroblasts (Driesen et al. 2005), but the functional characteristics of the link between cardiac myocyte and myofibroblast was not investigated. Connexin 45 has been identified within the MI scar of sheep hearts (Camelliti et al. 2004a), while in rabbit atria, Cx40 and Cx45 were found (Camelliti et al. 2004b). It is possible that these isoforms of connexin were also expressed in 24 h co-cultures of ventricular myofibroblasts and myocytes. However, incubating the cells with anti-Cx40 antibodies did not yield specific staining (data not shown), and an appropriate anti-Cx45 antibody is not commercially available. Partial cell fusion between fibroblasts and cardiomyocytes after 48 h in co-culture was reported recently (Driesen et al. 2005) and suggests an alternative mechanism for the intercellular communication observed in the present study.

The influence of ventricular myocyte activity on myofibroblast [Ca2+]i

When myofibroblasts were maintained for 24 h in co-culture with myocytes, stimulation of the myocyte was associated with an increase in [Ca2+]i in surrounding myofibroblasts. Such coupling of [Ca2+]i between co-cultured adult ventricular myocytes and myofibroblasts or fibroblasts has not previously been described. While the kinetics of the Ca2+ transients in the ventricular myocyte were rapid, the rise of [Ca2+]i in the myofibroblasts was relatively slow. The mechanism underlying changes in myofibroblast [Ca2+]i is not clear; one possibility is that stimulation of the myocyte induced the release of humoral factors that acted on the myofibroblasts in a paracrine manner. One candidate is ATP, a factor released from ventricular myocytes (Vassort, 2001) and known to initiate Ca2+ release in myofibroblasts (Brilla et al. 1998; Meszaros et al. 2000). The kinetics of the Ca2+ response to exogenous application of a high concentration of ATP were much more rapid than those of the response to ventricular myocyte stimulation, indicating that release of ATP from the ventricular myocyte would have to have a very slow time course to account for the responses observed in the myofibroblast. However, it is unlikely that this is the explanation, since after 4 h co-culture, paracrine interaction would be possible, yet field stimulation of myocytes was not associated with a change in myofibroblast [Ca2+]i. An alternative explanation lies in the presence of myocyte–myofibroblast coupling. Intercellular coupling may initiate Ca2+ changes either by the stimulation-dependant rise of [Ca2+]i that spreads from the myocyte into the surrounding myofibroblasts, similar to the spread of calcein dye, or by electrotonic coupling, which may trigger Ca2+ influx via changes in myofibroblast membrane potential. The latter possibility is unlikely because no significant inward Ca2+ current was observed in electrophysiological studies of myofibroblasts (Chilton et al. 2005). Furthermore, when applying voltage steps from −90 to 30 mV to myofibroblasts, the fura-2 ratio was unaffected (Fig. 7), indicating that the voltage step was not associated with a change in [Ca2+]i. The Na+–Ca2+ exchanger (NCX) is another route for Ca2+ entry in depolarized cells, but there is evidence to suggests that myofibroblasts may not express NCX (Koban et al. 2001). Alternatively, the connexin-based intercellular connections between myocytes and fibroblasts observed in this study may provide the pathway for Ca2+ exchange between myocyte and fibroblast. In support of this, heptanol, a compound known to disrupt gap junction function, abolished the changes in fibroblast [Ca2+]. However, further work is required to isolate the cellular basis of the rise of myofibroblast [Ca2+]i. The role of the rise of myofibroblast [Ca2+]i is unknown, but one possibility is that raised [Ca2+]i will increase contractile activity within the myofibroblasts, which may be mechanically beneficial in the context of an infarct scar.

Effect of intercellular coupling with myofibroblasts on myocyte electrophysiology

In the present study, intercellular coupling between ventricular myocytes and myofibroblasts did not appear to significantly affect myocyte electrophysiology unless a quasi-linear current was induced in the myofibroblast by stimulation with S-1-P. Action potential parameters were not statistically different between ventricular myocytes maintained under mono- versus co-culture conditions (Table 1). It is possible that subtle differences existed between these two groups, but the variability within each group obscured these differences. For example, judging from the size of the standard errors in the APD values, it is possible that an effect of co-culture of ±25 ms would be hidden by the variability within the data sets. This would suggest that the number of myofibroblasts directly coupled to the myocyte was small, thereby presenting a small additional current drain on the myocyte in the absence of S-1-P.

Effect of S-1-P on myofibroblast and myocyte electrophysiology

Sphingosine-1-phosphate activated a quasi-linear current in the myofibroblast at concentrations too low to directly affect the ventricular myocytes. Activation of this current increased the inward current required to hold the myofibroblast at −90 mV by approximately sevenfold and was associated with loss of excitability in co-cultured ventricular myocytes. The two events could be linked, in that the abrupt depolarization of the myocyte could be caused by a gradual increase in depolarizing currents in coupled myofibroblasts, eventually reaching the threshold required to abruptly depolarize the myocyte as illustrated in Fig. 8. Support for this hypothesis comes from the blockade of the effects of S-1-P on ventricular myocyte electrophysiology by the gap junction blocker heptanol.

The identity of the S-1-P-sensitive ion channel in adult rabbit cardiac myofibroblasts is unknown. Sphingosine-1-phosphate acts at G-protein coupled receptors of the S-1-P or Endothelial differentiation gene (EDG) family, which are expressed in the heart (Alewijnse et al. 2004). In guinea-pig atrial myocytes, S-1-P (1–100 nmol l−1) activated the acetylcholine-sensitive K+ current (IK,ACh (Ochi et al. 2006). In rabbit atrial myocytes, 1 μmol l−1 S-1-P activated IK,ACh as well. In contrast, the present study did not find a change in ventricular myocyte electrophysiology following 2 μmol l−1 S-1-P application, indicating that ventricular myocytes are relatively insensitive to S-1-P. In rabbit atrial myocytes, S-1-P (1 μmol l−1) antagonized the effect of isoproterenol on both L-type Ca2+ current (ICa,L) and hyperpolarization-activated inward current (If; Guo et al. 1999). Sphingosine-1-phosphate (3 μmol l−1) also inhibited ventricular myocyte Na+ current and depressed myocyte excitability (MacDonell et al. 1998). The present study is the first to show S-1-P-induced currents in cardiac myofibroblasts at considerably lower concentrations than those used previously in myocyte studies. In cultured fetal human lung myofibroblasts, S-1-P activated a Cl current (Yin & Watsky, 2006) with a similar current–voltage relationship to the current recorded in this study. Further studies are required to identify the ion channels activated by S-1-P in adult rabbit ventricular myofibroblasts.


Our results demonstrate that, following 24 h in co-culture, adult rabbit ventricular myocytes and myofibroblasts form intercellular junctions. The observed coupling of [Ca2+]i levels between activated ventricular myocytes and myofibroblasts may have implications for Ca2+-dependent processes within the myofibroblasts, such as scar contraction, secretion of autocrine/paracrine hormones, or collagen formation. In contrast, while this electrical coupling did not appear to significantly influence myocyte excitability or action potential characteristics under control conditions, activation of a quasi-linear conductance in myofibroblasts following S-1-P treatment caused marked depolarization of electrically coupled ventricular myocytes and loss of excitability.



The work was funded by the British Heart Foundation. The authors are indebted to Dr Glen Chilton, Dr Francis Burton, Dr Robert Clark and Professor Peter Kohl for their thoughtful comments. L.C. held a Postdoctoral Fellowship from the British Heart Foundation and was supported by the Alberta Heritage Medical Research Foundation (AHMRF) and the Heart and Stroke Foundation of Canada (HSF). W.R.G. is supported by the Canadian Institutes of Health and Research, the AHFMR, the HSF and funding from the University of California San Diego.