The electrotonic architecture of the retinal microvasculature: modulation by angiotensin II

Non-technical summary

In the quest to understand how the circulatory system adjusts microvascular function to meet local metabolic demand, we focused on the retina whose circulatory system consists exclusively of microvessels. Since voltages induced by extracellular signals play a key role in generating vasomotor responses, we characterized the movement of voltage within the retinal microvasculature. To do this, we quantified voltage transmission between pairs of recording pipettes located at well-defined sites in capillary/arteriole plexuses freshly isolated from the rat retina. We found that the retinal microvasculature is not simply a homogeneous syncytium, but has a complex electrotonic architecture with differing efficacies of voltage transmission. Furthermore, we discovered that the electrotonic architecture is not static, but is modulated by angiotensin. This newly appreciated action reveals that vasoactive signals can alter the functional organization of the microvasculature and, thereby, regulate the spatial extent of the circulatory system's response to voltage-changing inputs.

Abstract

Abstract

The capillary/arteriole complex is the key operational unit regulating local perfusion to meet metabolic demand. However, much remains to be learned about how this multicellular unit is functionally organized. To help address this challenge, we characterized the electrotonic architecture of the retinal microvasculature, which is particularly well adapted for the decentralized control of blood flow. In this study, we quantified the transmission of voltage between pairs of perforated-patch pipettes sealed onto abluminal cells located on microvascular complexes freshly isolated from the adult rat retina. These complexes consisted of capillaries, as well as tertiary and secondary arterioles. Dual recording experiments revealed that voltage spreading axially through a capillary, tertiary arteriole or secondary arteriole is transmitted very efficiently with a decay rate of only ∼5% per 100 μm. However, the retinal microvasculature is not simply a well-coupled syncytium since we detected significant voltage dissipation with radial abluminal cell-to-endothelium transmission and also at branch points between a capillary and its tertiary arteriole and between tertiary and secondary arterioles. Consistent with capillaries being particularly well-suited for the task of transmitting voltages induced by vasoactive signals, radial transmission is most efficient in this portion of the retinal microvasculature. Dual recordings also revealed that angiotensin II potently inhibits axial transmission. As a functional consequence, the geographical extent of the microvasculature's response to voltage-changing inputs is markedly restricted in the presence of angiotensin. In addition, this effect of angiotensin established that the electrotonic architecture of the retinal microvasculature is not static, but rather, is dynamically modulated by vasoactive signals.

Introduction

Capillaries may play a more active role in the regulation of blood flow than traditionally thought (Beach et al. 1998; McGahren et al. 1998; Schonfelder et al. 1998; Peppiatt et al. 2006; Puro, 2007). Significant progress in establishing this previously unappreciated function for capillaries has come from studies of the retinal vasculature. In the quest to better understand how local perfusion is regulated, investigators have focused on the circulatory system of the retina since it appears to be particularly well adapted for the decentralized control of blood flow. Indicative of this adaptation, retinal capillaries possess a high density of abluminally positioned pericytes (Shepro & Morel, 1993), which have a number of myocyte-like characteristics (Joyce et al. 1985; Tilton, 1991; Shepro & Morel, 1993; Hirschi & D'Amore, 1996) and whose contractions alter the capillary lumen (Schonfelder et al. 1998; Kawamura et al. 2003; Peppiatt et al. 2006; Puro, 2007) and thereby may affect local perfusion.

Evidence is accumulating that the retinal microvasculature is an interactive complex that includes a network of capillaries and a tertiary arteriole that links the capillaries with a secondary arteriole. Consistent with this operational concept, gap junction pathways link retinal capillaries to the proximal microvasculature (Oku et al. 2001). As in the microvasculature of other tissues (Song & Tyml, 1993; McGahren et al. 1998; Cohen et al. 2000), a functional manifestation of this interconnectivity is the observation that voltages generated in the capillaries of the retina spread efficiently to proximal sites (Ishizaki et al. 2009). Furthermore, localized electrical stimulation of a pericyte in the intact retina not only increases the contractile tone of the stimulated pericyte, but also evokes contractions in distantly located abluminal cells (Peppiatt et al. 2006).

Indicative of the complex operational organization of the retinal microvasculature, recent studies have revealed functional sub-specialization within this complex. For example, functional K_ATP channels are predominately located in the capillaries (Ishizaki et al. 2009) while functional voltage-dependent calcium channels (VDCCs) are chiefly in the proximal portions of the microvasculature (Matsushita et al. 2010). As a consequence, in order to initiate a VDCC-driven vasomotor response, a K_ATP-mediated voltage change generated in the capillaries must be transmitted proximally via gap junction pathways (Matsushita et al. 2010).

Despite recent progress in elucidating the organizational complexity and interactivity of the retinal microvasculature, much remains to be learned. To help address this gap in knowledge, a goal of this study was to quantitatively characterize the electrotonic architecture of the capillary–tertiary arteriole–secondary arteriole complex. Another goal was to determine whether the electrotonic architecture of the retinal microvasculature is static or whether it is subject to modulation by extracellular signals, such as angiotensin II. This molecule was of interest because we previously observed that exposure of retinal capillaries to angiotensin not only causes pericytes to contract as stored calcium is released and calcium-permeable non-specific cation channels are activated, but also results in a decrease in the membrane capacitance (C_m), which may reflect a lessening of cell coupling (Kawamura et al. 2004). However, because C_m can be affected by membrane changes unrelated to cell coupling (Gillis, 1995), this study sought to definitively determine whether angiotensin modulates electrotonic transmission within the retinal microvasculature.

To characterize the electrotonic architecture of the retinal microvasculature, we quantified the spread of depolarization between pairs of perforated-patch pipettes sealed onto abluminal cells located in microvascular complexes freshly isolated from the adult rat retina. These complexes included a secondary arteriole, a tertiary arteriole and a network of capillaries (Matsushita & Puro, 2006; Ishizaki et al. 2009). An experimental advantage of this preparation is that it allows analysis of the intact capillary/arteriole complex, rather than simply capillary fragments as we studied previously (Wu et al. 2006). Also, it is easy to visually identify the abluminal cells of each portion of the retinal microvasculature, i.e. the pericytes of the capillaries, the myocytes of the tertiary arterioles and the encircling smooth muscle cells of the secondary arterioles (Matsushita & Puro, 2006). Thus, with isolated microvessels, it is relatively straightforward to quantify electrotonic transmission between a pair of perforated-patch pipettes that are sealed onto abluminal cells at two well defined locations within the retinal microvasculature.

In this first characterization of the electrotonic architecture of the retinal microvasculature, we found that axial transmission is highly efficient within the capillaries, tertiary arterioles and secondary arterioles. However, a significant dissipation of voltage occurs at the branch point of a capillary and a tertiary arteriole, as well as at the tertiary arteriole–secondary arteriole junction. In addition, our experiments showed that voltage dissipation also occurs with radial transmission between an abluminal cell and the endothelium. Consistent with the capillary network being particularly well-suited for generating and transmitting voltages, radial transmission is most efficient in this portion of the retinal microvasculature. Importantly, our observation that angiotensin II selectively and profoundly inhibits axial transmission lends support for the operational concept that the electrotonic architecture of the retinal microvasculature is not static, but rather, is dynamically modulated by vasoactive signals.

Methods

Animal use conformed to the guidelines set forth by the Association for Research in Vision and Ophthalmology and was approved by the University of Michigan Committee on the Use and Care of Animals. The experimental procedures also comply with the polices set out by The Journal of Physiology (Drummond, 2009). This study used 76 Long–Evans rats (Charles River, Cambridge, MA, USA), which were 6–9 weeks old, weighed between 125 and 325 g and included approximately equal numbers of males and females. The animals were maintained on a 12 h alternating light–dark cycle and received food and water ad libitum. Death was induced with a rising concentration of carbon dioxide.

Microvessel isolation

A tissue print technique (Ishizaki et al. 2009) was used to isolate microvascular complexes from the retinas of male and female rats that were 6–9 weeks old. In brief, the procedure for microvessel isolation included the rapid removal of retinas, excision of adherent vitreous, and incubation for ∼24 min at 30°C in Earle's balanced salt solution supplemented with 0.5 mm EDTA, 6 U papain (Worthington Biochemical Corp., Lakewood, NJ, USA) and 2 mm cysteine. Subsequently, retinas were placed in solution A and quadrisected, where solution A consisted of 140 mm NaCl, 3 mm KCl, 1.8 mm CaCl₂, 0.8 mm MgCl₂, 10 mm Na-Hepes, 15 mm mannitol, and 5 mm glucose at pH 7.4 with osmolarity adjusted to 310 mosmol l⁻¹. Each quadrant of retina was then positioned vitreal-surface-up onto the glass bottom of a chamber containing solution A and gently compressed by a glass coverslip (15 mm diameter, 0.15 mm thick; CS-15R, Warner Instrument Corp., Hamden, CT, USA) onto which microvascular complexes adhered. As shown in published photomicrographs (Matsushita & Puro, 2006; Ishizaki et al. 2009) and the schematic diagram in Fig. 1, the isolated microvascular complexes used in this study included, from proximal to distal, a secondary arteriole encircled by a single layer of ‘doughnut-shaped’ smooth muscle cells, a tertiary arteriole with a layer of ‘dome-shaped’ myocytes at a density of ≥5 somas per 100 μm, and a capillary network whose abluminal cells, the pericytes, appear as ‘bumps on a log’ (Kuwabara & Cogan, 1960) with a density of ≤4 per 100 μm. As documented previously (Ishizaki et al. 2009), approximately 200 μm from its junction with a myocyte-encircled secondary arteriole, a tertiary arteriole bifurcates into two branches each of which typically extends another ∼200 μm before splitting into a pair of capillaries.

Figure 1. Schematic diagram showing the portion of the rat retinal microvasculature isolated by the tissue print procedure used in this study.

In this study, perforated-patch pipettes were sealed onto smooth muscle cells that encircle secondary arterioles, myocytes that are positioned on the endothelium of tertiary arterioles and abluminal pericytes located on the capillaries. Modified from Matsushita et al. 2010 with permission from the Association for Research in Vision and Ophthalmology.

Electrophysiology

Experiments were performed at room temperature, i.e. 22–23°C, within 5 h after microvessel isolation. In some experiments, a micromanipulator-guided micropipette was used to transect freshly isolated microvascular complexes at the capillary–secondary arteriole and the tertiary arteriole–secondary arteriole junctions. Details of this transaction technique are available (Ishizaki et al. 2009).

Perforated-patch recordings were made at sites along a freshly isolated microvessel located on a coverslip positioned in a recording chamber whose perfusate was solution A without or with 500 nm angiotensin II. The solution filling the recording pipettes consisted of 50 mm KCl, 65 mm K₂SO₄, 6 mm MgCl₂, 10 mm K-Hepes, 60 μg ml⁻¹ amphotericin B and 60 μg ml⁻¹ nystatin at pH 7.35 and osmolarity 280 mosmol l⁻¹. Recording pipettes had resistances of ∼5 MΩ and were mounted in the holder of a patch-clamp amplifier (Axopatch 200B, Molecular Devices, Sunnyvale, CA, USA or Dagan 3900, Dagan Corp., Minneapolis, MN, USA). The tip of a recording pipette was sealed with a resistance of ≥10 GΩ onto an abluminal cell, i.e. a smooth muscle cell encircling a secondary arteriole, a myocyte positioned on the wall of tertiary arteriole or a pericyte located on a capillary. In microvascular complexes isolated by our tissue print technique, it is straightforward to visually identify these abluminal cells (Matsushita & Puro, 2006). Of note, to confirm that recording pipettes could reliably be sealed onto pericytes and to assess gap junction-mediated interactions of these abluminal cells, we previously (Oku et al. 2001; Kawamura et al. 2002) used standard whole-cell pipettes loaded with Neurobiotin, which is a gap junction-permeant tracer. Under conditions that inhibit gap junction function, we found that 36 of 36 recording pipettes that were judged by visual inspection to be sealed onto a capillary pericyte were in fact so located since only a single pericyte became loaded with Neurobiotin diffusing from the pipette (Oku et al. 2001); endothelial cells were never labelled under conditions in which gap junctions were closed. Our studies also indicated that under control conditions, Neurobiotin spreads from a sampled pericyte to the underlying endothelium (Oku et al. 2001).

The access resistance for the recordings used in this study was <25 MΩ. Currents and voltages were filtered with a four-pole Bessel filter, sampled digitally at 500 or 2000 Hz using a DigiData 1440A acquisition system (Molecular Devices) and stored by a computer equipped with pCLAMP (version 10, Molecular Devices), which along with other software (Origin, v. 8.1, OriginLab Corp., Northampton, MA, USA), aided with data analysis and graphics display. Junction potential adjustment was made after data collection. The resting membrane potentials recorded from the capillaries, tertiary arterioles and secondary arterioles of isolated retinal microvascular complexes were –43 ± 1 mV (n = 27), –42 ± 1 mV (n = 29) and –45 ± 1 mV (n = 10), respectively; these potential were not significantly different. The input resistance calculated using the change in current induced by a 20 mV hyperpolarization from a holding potential of −58 mV was 175 ± 19 MΩ, 151 ± 9 MΩ and 127 ± 15 MΩ for voltage-clamp recordings from capillaries, tertiary arterioles and secondary arterioles, respectively; these resistances were not significantly different.

Dual recording experiments

Dual perforated-patch recordings provided a measure the efficacy of electrotonic transmission within the retinal microvasculature. In these experiments, voltages were monitored via pipettes sealed onto abluminal cells located at two sites along a microvessel while a 750 ms step of current was injected at 3 s intervals via one of the pipettes. For each current step, the ratio of the voltage change detected at the non-stimulated site (ΔV_responder) to the voltage step induced at the site of current injection (ΔV_stimulator) was calculated. Each value plotted in Fig. 3 is the mean of at least 30 successive ΔV_responder/ΔV_stimulator ratios. Because the effect of angiotensin on the ΔV_responder/ΔV_stimulator ratio was transient, the values plotted in Fig. 7 are the average of the three successive ratios obtained during the maximal effect of angiotensin on this ratio. First-order exponential fits of the dual recording data presented in Figs 3 and 7 were made using commercially available software (Origin, v. 8.1). A photomicrograph of the each studied microvascular complex provided documentation of the location of each recording pipette in the retinal microvasculature and aided in the determination of the distance between the pair of recording pipettes.

Figure 3. Plots of ΔV_responder/ΔV_stimulator ratios versus the distance between a pair of recording sites.

For each data set, the first-order exponential fit is shown. A, ratios for recordings from pairs of pericytes located on capillaries. B, ratios for dual recordings from pairs of myocytes located on tertiary arterioles. C, ratios for dual recordings from pairs of smooth muscle cells that encircled secondary arterioles. The mean of the ratios in panel A (0.56 ± 0.03) was significantly (P < 0.001) different than those in panels B (0.27 ± 0.02) and C (0.25 ± 0.06); the ratios in B and C are not significantly different.

Figure 7. ΔV_responder/ΔV_stimulator ratios versus the distance between a pair of recording sites on microvessels during exposure to 500 nm angiotensin.

A, ratios for recordings from pairs of pericytes located on capillaries. B, ratios for dual recordings from pairs of myocytes located on tertiary arterioles. C, ratios for dual recordings from pairs of smooth muscle cells located on secondary arterioles. For each panel, the first-order exponential fit is shown. Also shown in each panel is the first-order exponential fit for the ΔV_responder/ΔV_stimulator ratios obtained under control conditions and plotted in Fig. 3.

Of note, conduction through the bathing solution did not contribute to the observed ΔV_responder since pipette-to-pipette transmission was not detected after one of the two recording seals was spontaneously lost or when one of the pipettes was positioned close to, but not sealed onto, the microvessel. As we observed previously (Wu et al. 2006), there was no significant difference in the ΔV_responder/ΔV_stimulator ratios generated by a depolarization or a hyperpolarization; neither did the direction of transmission, i.e. distal to proximal or proximal to distal, significantly affect this ratio.

To calculate the velocity at which a voltage was conducted between two recording sites, the interpipette distance was determined from a photomicrograph of the sampled microvessel and with the aid of pCLAMP software, the time interval between the onset of the depolarization induced in the current-injected cell and the onset of the depolarization detected at the distant recording site was determined. For these calculations, dual recordings with interpipette distances of ≥200 μm were used.

Single perforated-patch recordings

In some experiments (Figs 10 and 11), voltage steps from a holding potential of –58 mV were used to generate current–voltage plots. As done previously (Sakagami et al. 1999; Kawamura et al. 2003; Kawamura et al. 2004), the amplitude of the non-specific cation current was measured from recordings in which the voltage was stepped to –103 mV, which is the K⁺ equilibrium potential. For the calculation of conductance densities, membrane capacitances were determined by the method of Zhao & Santos-Sacchi (1998).

Figure 10. Effect of angiotensin on current–voltage relations recorded from capillaries, tertiary arterioles and secondary arterioles.

A, averaged I–V plots generated from single perforated-patch recordings made from 8 pericyte located on capillaries whose connections with tertiary arterioles had been transected. ○, before angiotensin; ▾, during exposure to 500 nm angiotensin. At each tested voltage from –38 mV to –108 mV, the inward current was significantly (P < 0.0001) larger during exposure to angiotensin; at 12 mV and 22 mV, the outward current during angiotensin exposure was significantly (P < 0.05) larger. B, averaged I–V plots generated from perforated-patch recordings made from 6 myocytes located on tertiary arterioles whose connections with capillaries and secondary arterioles had been transected. ○, before angiotensin; ▾, during exposure to 500 nm angiotensin. Angiotensin did not significantly affect the current amplitude. C, averaged I–V plots generated from perforated-patch recordings made from 3 smooth muscle cells located on secondary arterioles whose connections with tertiary arterioles had been transected. ○, before angiotensin; ▾, during exposure to 500 nm angiotensin. Angiotensin did not significantly affect the current amplitude.

Figure 11. Angiotensin-induced non-specific cation (NSC) conductance density in capillaries, tertiary arterioles and secondary arterioles, which were part of an intact microvascular complex or that had been transected from the rest of the microvascular plexus.

For the capillary groups, 8 recordings were from pericytes located on transected capillaries, and 6 were from pericytes on capillaries of intact microvascular complexes. For the tertiary arteriole groups, 6 recordings were from myocytes located on transected tertiary arterioles, and 5 were from myocytes on tertiary arterioles of intact microvascular complexes. For the secondary arteriole groups, 3 recordings were from smooth muscle cells encircling transected secondary arterioles, and 4 were from smooth muscle cells on secondary arterioles of intact microvascular complexes. For each portion of the microvasculature, the angiotensin-induced non-specific cation conductance density was not significantly different in intact and transected microvessels.

Immunocytochemistry

Unless otherwise noted, this protocol was performed at room temperature. After microvessel-containing coverslips were rinsed in phosphate-buffered saline (PBS) and fixed with 4% formaldehyde in PBS for 30 min, endogenous peroxidase activity was blocked by 0.3% hydrogen peroxide in PBS for 30 min. Coverslips were then exposed overnight at 4°C to a well-characterized primary anti-angiotensin receptor type 1 (AT₁) antibody (product no. AAR-011, Alomone Labs, Jerusalem, Israel) that was diluted 1:100 in PBS supplemented with 1.5% normal goat serum. Two negative controls were used. In one control, the primary antibody was omitted. As a second negative control, the primary antibody was pre-incubated with the antigen peptide (NSSTEDGIKRIQDDC, which corresponds to amino acid residues 4–18 of human AT₁ receptor) for 1 h preceding application onto microvessels. After incubation with biotin-conjugated goat anti-rabbit IgG (1:200, Vector Laboratories, Inc., Burlingame, CA, USA) for 1 h, coverslips were kept for ∼40 h at 4°C in a horseradish peroxidase-streptavidin solution (RTU, Vector Laboratories) and then exposed to the avidin–biotin–peroxidase complex (1:100, ABC method, Vector Laboratories) for 30 min. After development of diaminobenzidine (DAB kit, Vector Laboratories), photomicrographs were taken with differential interference contrast optics.

Chemicals

Unless otherwise noted, chemicals were obtained from Sigma-Aldrich (St Louis, MO, USA).

Statistics and data analysis

Data are given as means ± SEM. Probability was evaluated by Student's two-tailed t test, unless noted otherwise. For the comparison of two groups, P≥ 0.05 indicated lack of a significant difference. For the statistical comparison of three groups, the P-value was adjusted using the Bonferroni correction.

Results

Dual recordings from capillaries, tertiary arterioles and secondary arterioles

In order to characterize the electrotonic architecture of the retinal microvasculature, we quantified the spread of depolarization between pairs of perforated-patch pipettes sealed onto abluminal cells located on microvascular complexes freshly isolated from the adult rat retina. Each complex studied contained a capillary network branching from a tertiary arteriole that originated from a secondary arteriole (Fig. 1). An example of such an experiment is shown in Fig. 2. In the illustrated experiment, a pair of pipettes was sealed onto myocytes located on a tertiary arteriole (Fig. 2A and B). The experimental protocol was to inject a depolarizing current into one of the monitored myocytes while the change in voltage was monitored at both recording sites (Fig. 2C). The ratio of the voltage change measured at the non-stimulated site, i.e. ΔV_responder, to the voltage change in the current-injected cell, i.e. ΔV_stimulator, yielded a measure of the efficacy of electrotonic transmission between the pair of recording pipettes.

Figure 2. Example of a dual perforated-patch recording used to measure electrotonic transmission within a tertiary arteriole.

A and B, photomicrographs of the sampled microvessel showing the recording sites on a tertiary arteriole. Scale bars: 50 μm. C, left, voltage trace recorded by a pipette sealed onto an abluminal myocyte into which a depolarizing current was injected. Right, the voltage trace recorded via the pipette sealed onto myocyte located on the tertiary at a distance of 200 μm from the stimulated (current-injected) myoctye. For this pair of recordings from tertiary arteriolar myocytes, the ΔV_responder/ΔV_stimulator ratio was 0.34.

In a series of dual recording experiments, the ΔV_responder/ΔV_stimulator ratios were determined for a range of interpipette distances within capillaries, tertiary arterioles and secondary arterioles (Fig. 3). As is evident in Fig. 3, the interpipette distance had only a minimal effect on the ΔV_responder/ΔV_stimulator ratio. More specifically, first-order exponential fits of these data yielded voltage decay rates of 2 ± 2%/100 μm (n = 27) in the capillaries (Fig. 3A), 6 ± 8%/100 μm (n = 22) in the tertiary arterioles (Fig. 3B) and 6 ± 8%/100 μm (n = 10) in the secondary arterioles (Fig. 3C); these decay rates were not significantly different. Based on these findings, we concluded that axial transmission is highly efficient within the capillaries, the tertiary arterioles and the secondary arterioles of the retinal microvasculature.

Models of the electrotonic architecture of the retinal microvasculature

What is the intercellular pathway by which voltage is transmitted from a current-injected abluminal cell to another abluminal cell? One possibility considered was that electrotonic transmission is predominantly from abluminal cell-to-abluminal cell. Since the decay of voltage spreading along this homocellular pathway would be expected to be, at least approximately, a first-order process, the extrapolated ΔV_responder/ΔV_stimulator ratio at an interpipette distance of 0 μm should be 1.0. However, in contrast to this expectation, the first-order fits of the observed ΔV_responder/ΔV_stimulator ratios shown in Fig. 3 did not extrapolate to 1.0 at 0 μm. Rather, the extrapolated ratios at 0 μm were 0.58 ± 0.07, 0.29 ± 0.03 and 0.22 ± 0.04 for capillaries, tertiary arterioles and secondary arterioles, respectively (Fig. 3). Thus, this analysis of the ΔV_responder/ΔV_stimulator data in Fig. 3 indicates that direct abluminal cell-to-abluminal cell transmission is not likely to be the predominate pathway for the spread of voltage from a current-injected abluminal cell.

To further assess putative models for the spread of voltage through the retinal microvasculature (Fig. 4), the ΔV_responder/ΔV_stimulator ratios observed in recordings from adjacent abluminal cells, i.e. recordings with interpipette distances of ≤20 μm (Fig. 3), were initially used to calculate the decay of voltage in a model in which axial transmission was chiefly from abluminal cell to abluminal cell. In Fig. 4A and B, the observed ΔV_responder/ΔV_stimulator ratios are compared with the ratios calculated for homocellular abluminal cell-to-abluminal cell transmission. As shown in Fig. 4B, the homocellular model predicted that the ΔV_responder/ΔV_stimulator ratio at an interpipette distance of 200 μm would be <0.00. However, this predicted ratio did not accurately match the observed ratio of 0.26 (Fig. 4A). Similarly, we found that this homocellular model failed to account for the ΔV_responder/ΔV_stimulator ratios observed for distantly separated recording pairs in capillaries and secondary arterioles.

Figure 4. Models of the electrotonic architecture.

Each schematic drawing shows a tertiary arteriole with four successively located abluminal myoctyes, as well as a myocyte located 200 μm from the current-injected myocyte. A relative voltage change of 1.0 was generated in the abluminal cell located on the extreme left side. Of note, in tertiary arterioles of retinal microvascular complexes, there are ≥5 myocytes per 100 μm (Matsushita & Puro, 2006). A, relative voltage changes observed in dual recordings from tertiary arterioles; data are from Fig. 3B. B, model of homocellular abluminal cell-to-abluminal cell transmission. Within each myocyte is shown the relative voltage change calculated using 0.31 as the efficacy of axial transmission between adjacent myoctyes; 0.31 was the ΔV_responder/ΔV_stimulator ratio observed in dual recordings from adjacent myocytes (Fig. 3B). Comparison of the calculated ratios shown in this panel with the observed ratios in panel A shows that this homocellular model failed to adequately predict the ΔV_responder/ΔV_stimulator ratio observed at an interpipette distance of 200 μm. C, heterocellular model for the spread of voltage from a current-injected abluminal cell. As detailed in the text and listed in Table 3, the experimentally derived transmission efficacies used to calculate the relative voltage changes in this heterocellular model were 0.54 for radial myocyte-to-endothelium transmission, 0.94/100 μm for axial transmission through the arteriolar endothelium and 0.54 for endothelium-to-myocyte transmission at the site of the distantly monitored myocyte. The ΔV_responder/ΔV_stimulator ratios calculated using this heterocellular model closely matched the observed ratios shown in panel A. D, relative voltage changes observed in dual recordings from tertiary arterioles during exposure to angiotensin; data are from Fig. 7B. E, heterocellular model in the presence of angiotensin. As detailed in the text and listed in Table 3, the experimentally derived transmission efficacies used to calculate the relative voltage changes were 0.59 for radial myocyte-to-endothelium transmission, 0.52/100 μm for axial transmission through the arteriolar endothelium and 0.59 for endothelium-to-myocyte transmission at the site of the distantly monitored myocyte; only axial transmission was significantly affected by angiotensin. The ΔV_responder/ΔV_stimulator ratios calculated using this heterocellular model closely matched the observed ratios shown in panel D.

What model can accurately account for the ΔV_responder/ΔV_stimulator data presented in Fig. 3? We considered a heterocellular model in which voltage spreads from a current-injected abluminal cell via the following pathway: radially from that abluminal cell to the underlying endothelium, then axially through the endothelial layer and finally radially from the endothelium to the distantly located abluminal cell (Fig. 4C). In addition, in order to account for substantial dissipation of voltage being detected in dual recordings made at short interpipette distances and only minimal additional decay with longer interpipette distances (Figs 3 and 4A), our heterocellular model has relatively inefficient radial transmission and highly efficient axial transmission.

Experimental evidence supports the heterocellular model. Consistent with there being radial communication between abluminal cells and the endothelium, we reported previously that loading the tracer Neurobiotin into abluminal cells via whole-cell recording pipettes resulted in the gap junction-dependent spread of this tracer into and extensively through the endothelial layer of the retinal microvasculature (Oku et al. 2001; Kawamura et al. 2002). Also consistent with the heterocellular model in which axial transmission through the endothelium is highly efficient, our dual recording experiments indicated that voltage spreading axially along a retinal microvessel decays only ∼5% per 100 μm (Fig. 3). Also consistent with our heterocellular model, dual recording experiments indicated that radial transmission is less efficient. Namely, a parsimonious explanation for the ΔV_responder/ΔV_stimulator ratios in Fig. 3 extrapolating to <1.0 at 0 μm is that the extrapolated ratio reflects the dissipation of voltage during radial abluminal cell/endothelium transmission at the current-injected site and also at the distantly recorded site. Thus, for capillaries, the extrapolated ΔV_responder/ΔV_stimulator ratio of 0.58 ± 0.07 at the interpipette distance of 0 μm (Fig. 3A) is the product of the transmission efficacy of 0.76 ± 0.05 for the passage of voltage from a current-injected pericyte to the underlying endothelium and 0.76 ± 0.05 for the passage of voltage from the endothelium to the distantly monitored pericyte, i.e. the overall transmission efficacy for these two radial transmission steps is 0.76 × 0.76 = 0.58. Similarly for tertiary arterioles, the extrapolated ΔV_responder/ΔV_stimulator ratio of 0.29 ± 0.03 at the interpipette distance of 0 μm (Fig. 3B) yields a radial transmission efficacy of 0.54 ± 0.04. For secondary arterioles, the extrapolated value of 0.22 ± 0.04 (Fig. 3C) yields a radial transmission efficacy of 0.47 ± 0.06. Of note, these efficacies for radial transmission in the arterioles were not significantly different. However, of likely functional importance, the efficacy of radial transmission in the capillaries was significantly (P = 0.0027) greater in the capillaries than in the arterioles.

As illustrated in Fig. 4C, our heterocellular model for the spread of voltage from a current-injected abluminal cell provided a reasonable prediction of the observed ΔV_responder/ΔV_stimulator ratios. Thus, even though we do not exclude that there may be some transmission along the abluminal cell layer, a heterocellular model in which radial abluminal cell–endothelium transmission and endothelial cell–endothelial cell transmission predominate appears to account well for the observed results of the dual recording experiments presented in Fig. 3.

Effect of branch points on electrotonic transmission

In addition to quantifying electrotonic transmission within non-branching segments of intact microvascular complexes, we also assessed the effect of branch points within the retinal microvascular complex. In a series of eight experiments, dual recordings were made in which one perforated-patch pipette was sealed onto a pericyte located on a capillary and the other pipette was sealed onto an abluminal myocyte of the tertiary arteriole. The ΔV_responder/ΔV_stimulator ratios observed in these experiments are shown in Table 1. Also shown in Table 1 are the ‘predicted’ ratios calculated using the efficacies of radial and axial transmission derived as detailed above. Consistent with a capillary/tertiary arteriole branch point diminishing the efficacy of transmission, the experimentally observed ΔV_responder/ΔV_stimulator ratio was significantly (P < 0.0001) less than the predicted ratio. To adjust the predicted ratio to match the observed ratio, the transmission efficacy at the capillary–tertiary arteriole junction was calculated to be 0.57 ± 0.03.

Table 1.

Dual recording experiments designed to determine the effect of the capillary–tertiary arteriole branch point on the efficacy of electrotonic transmission

Pair	Capillary distance (μm)	Tertiary arteriole distance (μm)	Observed ΔVresponder/ΔVstimulator	Predicted ΔVresponder/ΔVstimulator	Capillary-to-tertiary arteriole transmission efficacy
1	128	325	0.20	0.35	0.57
2	58	122	0.22	0.38	0.58
3	155	205	0.20	0.36	0.56
4	290	90	0.21	0.36	0.58
5	205	305	0.20	0.34	0.59
6	335	33	0.14	0.37	0.39
7	220	85	0.24	0.37	0.65
8	160	170	0.23	0.36	0.64
			0.20 ± 0.01	0.36 ± 0.004*	0.57 ± 0.03

For each of 8 dual recording experiments, one perforated-patch pipette was sealed onto a pericyte located on a capillary while the other recording pipette was sealed onto a myocyte located on the tertiary arteriole. The second and third columns show the lengths of the capillary and tertiary arteriole between the two recording sites. Also shown for each experiment is the observed ΔV_responder/ΔV_stimulator ratio and the ‘predicted’ ratio, which was calculated by using the transmission efficacies derived from the data in Fig. 3A and B, i.e. 0.76 for radial transmission between a pericyte and the underlying endothelium, 0.98/100 μm for axial transmission along the capillary, 0.94/100 μm for axial transmission along the tertiary arteriole, and 0.54 for transmission from the endothelium to the myocyte of the tertiary arteriole. For each pair of recordings, the observed ΔV_responder/ΔV_stimulator ratio was smaller than the predicted value; for this series of experiments, this was a significant difference (*P < 0.0001). The final column gives the calculated value that equalized the predicted ΔV_responder/ΔV_stimulator ratio with the experimentally determined ratio; this calculated value was deemed to be the efficacy of transmission at the capillary–tertiary arteriole branch point.

We also assessed the effect of branch points between tertiary and secondary arterioles. In series of seven dual perforated-patch recordings in which one pipette was sealed onto a myocyte of a tertiary arteriole and the other onto a smooth muscle cell encircling a secondary arteriole, the observed ΔV_responder/ΔV_stimulator ratio was significantly (P < 0.0001) less than that predicted by using the transmission efficacies determined for unbranched tertiary and secondary arterioles (Table 2). In order to account for the observed ΔV_responder/ΔV_stimulator ratios, the efficacy of transmission across the tertiary arteriole–secondary arteriole junction was calculated to be 0.62 ± 0.14, which was not significantly different from the transmission efficacy determined for the capillary/tertiary arteriole junction.

Table 2.

Dual recording experiments designed to determine the efficacy of electrotonic transmission at the tertiary arteriole–secondary arteriole junction

Cell	Tertiary arteriole distance (μm)	Secondary arteriole distance (μm)	Observed ΔVresponder/ΔVstimulator	Predicted ΔVresponder/ΔVstimulator	Tertiary arteriole-to-secondary arteriole transmission efficacy
1	22	61	0.12	0.25	0.48
2	42	58	0.14	0.25	0.56
3	33	63	0.16	0.25	0.64
4	5	55	0.17	0.25	0.68
5	50	13	0.22	0.25	0.88
6	50	270	0.10	0.22	0.45
7	60	270	0.15	0.22	0.68
			0.14 ± 0.02	0.24 ± 0.004*	0.62 ± 0.05

For each pair of recordings, one perforated-patch pipette was sealed onto a myocyte of a tertiary arteriole and the other onto a smooth muscle cell encircling a secondary arteriole. The second and third columns show the axial lengths of the tertiary arteriole and the secondary arteriole that were between the two recording sites. For each experiment, the table shows the observed ΔV_responder/ΔV_stimulator ratio and the ‘predicted’ ratio, which was calculated by using transmission efficacies derived from the data in Fig. 3B and C, i.e. 0.54 for radial transmission between the current-injected myocyte of a tertiary arteriole and the underlying endothelium, 0.94/100 μm for axial transmission along the endothelium of the tertiary arteriole, 0.94/100 μm for axial transmission along the endothelium of the secondary arteriole, and 0.47 for radial transmission from the endothelium to the smooth muscle cell monitored in the secondary arteriole. For each pair of recordings, the observed ΔV_responder/ΔV_stimulator ratio was smaller than the predicted value; for the series of experiments, this was a significant difference (*P < 0.0001). The final column gives the calculated value that equalized the predicted ΔV_responder/ΔV_stimulator ratio with the experimentally determined ratio; this value was deemed to be the efficacy of transmission at the site where a tertiary arteriole branches from a secondary arteriole.

In other dual perforated-patch recordings, we used analyses similar to those outlined above to assess the effects of branch points within tertiary arterioles and capillaries. We found that when one pipette was sealed onto a myocyte located on a branch of a tertiary arteriole and the other pipette was sealed onto a myocyte on the proximal stalk of this arteriole, the observed ΔV_responder/ΔV_stimulator ratio was 0.27 ± 0.03 (n = 16) at an interpipette distance of 307 ± 42 μm. Consistent with a branch point within a tertiary arteriole not affecting transmission, 0.27 was also the ΔV_responder/ΔV_stimulator ratio calculated by using the efficacies determined for radial and axial transmission within tertiary arterioles. Thus, it appears that branches within a tertiary arteriole did not cause voltage dissipation.

In other experiments, we recorded from pericytes at two capillary sites between which there was a branch. For this series of experiments, the observed ΔV_responder/ΔV_stimulator ratio at an interpipette distance of 236 ± 26 μm was 0.49 ± 0.06 (n = 6). Indicative that capillary branch points did not cause a spreading voltage to dissipate, 0.55 was the ratio calculated by using the efficacies of radial and axial transmission in the capillaries. From these data, we concluded that branches within capillaries, as well as those within the tertiary arterioles, did not affect electrotonic transmission. Thus, some classes of branch points, i.e. capillary–tertiary arteriole and tertiary arteriole–secondary arteriole, but not others affect the spread of voltage within the retinal microvasculature.

Transmission velocities

In addition to providing data to quantify electrotonic transmission, the voltage traces recorded from a current-injected abluminal cell and a distant responding abluminal cell permitted the calculation of the velocity at which a voltage is transmitted along a microvessel. We found that the conduction velocities along capillaries, tertiary arterioles and secondary arterioles were 50 ± 4 mm s⁻¹ (n = 8), 53 ± 2 mm s⁻¹ (n = 21) and 42 ± 10 mm s⁻¹ (n = 6), respectively; these velocities are not significantly different.

Taken together the results of our dual recording experiments support the idea that the electrotonic architecture of the retinal microvasculature is characterized chiefly by radial abluminal cell–endothelium transmission and axial transmission through the endothelial layer. Furthermore, the data indicate that radial transmission is less efficient than axial transmission within the capillaries, tertiary arterioles and secondary arterioles. In addition, we found that substantial voltage dissipation occurs at capillary–tertiary arteriole branch points and also at junctions of tertiary and secondary arterioles.

Effect of angiotensin on the electrotonic architecture

We considered the possibility that the electrotonic architecture of the retinal microvasculature is not static, but rather is dynamically modulated. Suggestive that the vasoactive signal, angiotensin II, may affect cell coupling, previous recordings from isolated retinal capillaries showed that activation of AT₁ receptors was associated with a decrease in membrane capacitance (Kawamura et al. 2004). However, because membrane capacitance provides, at best, only a qualitative suggestion of the extent of cell coupling, we used dual recordings to quantify the effect of angiotensin on electrotonic transmission. In addition, since immunoreactivity for AT₁ angiotensin receptors was found on abluminal cells and the endothelium throughout the retinal microvasculature (Fig. 5), this study assessed the effect of angiotensin on electrotonic transmission in secondary and tertiary arterioles, as well as in the capillary network.

Figure 5. Immunoreactivity for the AT₁ angiotensin II receptor.

A, photomicrograph showing a portion of a microvascular complex freshly isolated from the rat retina and stained with anti-AT₁ antibody. Microvascular cells throughout the retinal microvasculature were immunopositive. B, negative control in which a microvascular complex was prepared for immunocytochemistry in the absence of the primary antibody. C, negative control in which the primary antibody was pre-incubated with the antigenic peptide before exposure of a microvascular complex to the antibody/antigenic peptide-containing solution. Scale bars: 50 μm.

As illustrated in Fig. 6, dual perforated-patch recordings demonstrated that the ΔV_responder/ΔV_stimulator ratio transiently decreased during exposure to angiotensin. Using a similar protocol, ΔV_responder/ΔV_stimulator ratios at various interpipette distances were obtained in recordings from pairs of capillary pericytes (Fig. 7A), of tertiary arteriolar myocytes (Fig. 7B) and of secondary arteriolar smooth muscle cells (Fig. 7C). Analysis of the data in Fig. 7 showed that angiotensin caused the efficacy of axial transmission (Fig. 8A) to decrease significantly (P < 0.0001) from 0.98 ± 0.03/100 μm to 0.50 ± 0.05/100 μm (n = 6) in the capillaries, from 0.94 ± 0.08/100 μm to 0.52 ± 0.07/100 μm (n = 10) in the tertiary arterioles, and from 0.94 ± 0.08/100 μm to 0.55 ± 0.11/100 μm (n = 8) in the secondary arterioles. In contrast, because the extrapolated ΔV_responder/ΔV_stimulator ratios at 0 μm were not altered significantly by angiotensin (Fig. 7), it appears that radial transmission in capillaries, tertiary arterioles and secondary arterioles was unaffected by this vasoactive molecule (Fig. 8B). In addition, we did not detect a significant effect of angiotensin on transmission at the various branch points within the microvasculature. Thus, these experiments indicated that angiotensin selectively inhibited axial transmission.

Figure 6. Example of an experiment assessing the effect of angiotensin on electrotonic transmission.

A, photomicrograph showing the sites at which perforated-patch pipettes were sealed onto a capillary pericyte and the myocyte of tertiary arteriole. Scale bar: 50 μm. B, plot of the ΔV_responder/ΔV_stimulator ratio versus time. Each point is the mean of 3 successive sweeps. A depolarizing current was injected via the recording pipette that was sealed onto a capillary pericyte. Bar shows when 500 nm angiotensin was added to the perfusate. C, voltage traces from a current-injected capillary pericyte and from a myocyte on the tertiary arteriole when the bathing solution lacked angiotensin. D, voltage traces from the capillary pericyte and the myocyte on the secondary arteriole during exposure of the microvascular complex to angiotensin.

Figure 8. Effect of angiotensin on electrotonic transmission in the retinal microvasculature.

A, efficacies of axial transmission under control conditions and in the presence of 500 nm angiotensin. For each microvascular region, dual perforated-patch recordings were made from abluminal cells, i.e. pericytes of the capillaries, myocytes of the tertiary arterioles and encircling smooth muscle cells of the secondary arterioles. The number of dual perforated-patch experiments for each group is shown in Table 3. For each microvascular region, angiotensin significantly (*P < 0.0001) decreased the efficacy of axial transmission. B, efficacies of radial transmission under control conditions and in the presence of 500 nm angiotensin. Perforated-patch recordings were as described in panel A, and the number of dual perforated-patch experiments for each group is shown in Table 3. For each microvascular region, angiotensin did not significantly change the efficacy of radial transmission.

As shown in Fig. 4D and E, the observed ΔV_responder/ΔV_stimulator ratios obtained in the presence of angiotensin were accurately predicted by our heterocellular model when the efficacy of axial transmission was selectively decreased. Furthermore, as illustrated in Fig. 9, calculations that were based on our heterocellular model (Fig. 4C and E) and that used the efficacies of transmission derived from experiments performed under control conditions and during angiotensin exposure (Table 3) predicted that angiotensin would profoundly diminish the proximal spread of a voltage generated within the capillary tree.

Figure 9. Predicted voltage decay within the retinal microvasculature under control conditions and in the presence of angiotensin.

The vertical axis shows the relative voltage changes predicted to occur in abluminal cells during the spread of a relative voltage of 1.45 that was generated in the endothelium of a capillary at a site 400 μm distal to the capillary–tertiary arteriole junction. Relative voltages in abluminal cells were calculated at 100 μm intervals based on the heterocellular models illustrated in Fig. 4C and E and with the use of the efficacies of radial transmission, of axial transmission and of transmission at branch points listed in Table 3. Our heterocellular model of the electrotonic architecture of the retinal microvasculature predicts that the amount of a capillary-generated voltage spreading into and through the proximal arterioles is markedly decreased during exposure to angiotensin.

Table 3.

Efficacies of electrotonic transmission at various sites within the retinal microvasculature

Site within the retinal microvasculature	Efficacy of transmission under control conditions	Efficacy of transmission in angiotensin
Capillary
Axial transmission
Non-branching regions	0.98 ± 0.03/100 μm (n = 27)	0.50 ± 0.07/100 μm (n = 6)*
Branch points	1.12 ± 0.16 (n = 6)	1.11 ± 0.08 (n = 3)
Radial transmission	0.76 ± 0.05 (n = 27)	0.76 ± 0.05 (n = 6)
Capillary/tertiary arteriole branch point	0.57 ± 0.08 (n = 8)	0.62 ± 0.19 (n = 4)
Secondary arteriole
Axial transmission
Non-branching regions	0.94 ± 0.08/100 μm (n = 29)	0.52 ± 0.07/100 μm (n = 10)*
Branch points	1.00 ± 0.10 (n = 16)	1.14 ± 0.07(n = 3)
Radial transmission	0.54 ± 0.04 (n = 29)	0.59 ± 0.08 (n = 10)
Tertiary arteriole/secondary arteriole branch point	0.62 ± 0.05 (n = 7)	0.54 ± 0.12 (n = 4)
Secondary arteriole
Axial transmission
Non-branching regions	0.94 ± 0.08/100 μm (n = 10)	0.55 ± 0.11 (n = 8)*
Radial transmission	0.47 ± 0.06 (n = 10)	0.45 ± 0.13 (n = 8)

Values are derived from the data shown in Figs. 3 and 7 and in Tables 1 and 2. The n values are the number of pairs of perforated-patch recordings. Asterisks indicate that angiotensin significantly decreased the efficacy of axial transmission in the capillaries (P < 0.0001), the tertiary arterioles (P = 0.0057) and the secondary arterioles (P = 0.0097); at other microvascular sites, the transmission efficacy was not significantly affected by angiotensin.

Effect of angiotensin on the I–V relations of isolated capillaries and arterioles

In addition to quantifying the effect of angiotensin on the transmission of voltages generated by the injection of current, we wished to assess how angiotensin affected the spread of a physiological input. To do this, we used single perforated-patch recordings to measure the change in current–voltage relations induced by angiotensin, which is known to activate a non-specific cation conductance in the retinal microvasculature (Kawamura et al. 2004). For an initial series of experiments, microvascular complexes were transected at the capillary–tertiary arteriole junction and/or at the junction of the tertiary and secondary arterioles. In this way, we could obtain single perforated-patch recordings from abluminal cells located on isolated segments of capillaries, tertiary arterioles and secondary arterioles.

We found that during exposure to angiotensin, the transected capillaries generated a current that was inward at physiological membrane potentials and caused depolarization (Fig. 10A). In contrast, the current–voltage relations of transected tertiary arterioles and transected secondary arterioles were minimally affected by angiotensin (Fig. 10B and C). From these experiments, we concluded that the angiotensin-induced conductance is generated predominantly by the capillaries; little is generated by the arterioles. We also concluded from these observations that the lack a significant angiotensin-induced current in the arterioles indicates that a decrease in membrane resistance is not required in order for angiotensin to decrease the efficacy of axial transmission.

Effect of angiotensin on intact microvascular complexes

The finding that the angiotensin-induced current is generated predominately in the capillaries led to additional experiments designed to test the prediction of our heterocellular model (Fig. 9) that angiotensin's inhibition of axial transmission would minimize the spread of a capillary-generated current. In these experiments, single perforated-patch pipettes were sealed onto the capillaries, tertiary arterioles and secondary arterioles of intact microvascular complexes, and I–V relations were determined before and during exposure to angiotensin. As detailed in Methods, the angiotensin-induced NSC conductance was quantified at –103 mV. As summarized in Fig. 11, the angiotensin-induced conductances detected in the tertiary and secondary arterioles were not significantly different in recordings made from intact, as compared with transected, microvascular complexes. Thus, consistent with angiotensin potently attenuating axial transmission, essentially none of the angiotensin-induced current generated in the capillaries spread to the proximal microvasculature.

Discussion

To better understand the functional organization of the retinal microvasculature, we used dual perforated-patch recordings to characterize the electrotonic architecture of microvascular complexes freshly isolated from the adult rat retina. These experiments revealed that a voltage spreading axially along a capillary, a tertiary arteriole or a secondary arteriole is transmitted efficiently with a rate of decay of only ∼5% per 100 μm. However, our analysis also demonstrated that the retinal microvasculature is not simply a well-coupled syncytium. Rather, voltage is attenuated by ∼45% as it spreads from a branch of the capillary network to a tertiary arteriole and also as it passes from a tertiary arteriole to the proximal secondary arteriole. Our dual recording data also led us to infer that radial transmission between an abluminal cell and the underlying endothelium is not as efficient as axial transmission through the endothelium.

In this study, we also used dual recordings to assess the effect of angiotensin II on electrotonic transmission. These experiments indicated that this vasoactive molecule selectively inhibits axial transmission. Thus, the electrotonic architecture of the retinal microvasculature is not static, but rather can be dynamically modulated by vasoactive signals.

Comparison of retinal and non-retinal vasculatures

How does the electrotonic architecture of retinal microvasculature compare with the functional organization of other vascular systems? A key finding of our dual recording experiments is that the axial spread of voltage within retinal capillaries, tertiary arterioles and secondary arterioles is highly efficient. Although there appear to be no previous dual recording studies directly assessing electrotonic transmission within a capillary–arteriole complex, our finding of efficient axial transmission in retinal microvessels is consistent with studies demonstrating in non-retinal tissues that the application of voltage-changing chemicals onto a capillary can elicit vasomotor responses in proximal arterioles (Dietrich, 1989; Song & Tyml, 1993; Berg et al. 1997; Beach et al. 1998; McGahren et al. 1998). Furthermore, the voltage decay rate of ∼5%/100 μm within the retinal microvasculature is similar to the rate of decay for axial transmission through larger vessels in the circulatory system (Hirst & Neild, 1978; Emerson et al. 2002). In addition, our finding that a voltage spreads with a conduction velocity of ∼50 mm s⁻¹ along retinal capillaries, tertiary arterioles and secondary arterioles is similar to the velocities reported for larger vessels, such as those located in the ureter (23 mm s⁻¹) (Tsuchiya & Takein, 1990), retractor muscle (∼45 mm s⁻¹) (Emerson et al. 2002) and small intestine (∼85 mm s⁻¹) (Stevens et al. 2000). Thus, rapid and efficient axial transmission may be a feature, not only of the retinal microvasculature, but of the circulatory system in general.

In contrast to the apparent universality of highly efficient axial transmission, the efficacy of radial transmission varies widely. For example, in guinea-pig mesenteric secondary arterioles, the efficacy of transmission between an abluminal smooth muscle and the endothelium is only ∼6% (Yamamoto et al. 2001), while radial myoendothelial coupling is reported to be robust in arteries of the hamster retractor muscle (Emerson et al. 2002). Between these extremes, our study indicated the efficacy of radial transmission is 76% in the capillaries, 54% in the tertiary arterioles and 47% in the secondary arterioles (Table 3).

Another aspect of the electrotonic architecture that appears to vary within the circulatory system is the efficacy of transmission between abluminal cells. While transmission along the smooth muscle layer of hamster cheek pouch arteries is reported to be quite efficient (Bartlett & Segal, 2000; Budel et al. 2003), a current injected into a smooth muscle cell of a mesenteric secondary arteriole was found to dissipate by ∼90% after spreading just 25 μm along the abluminal layer (Yamamoto et al. 2001). Similarly, our model-based analysis of dual recording experiments indicates that abluminal cell–abluminal cell is, at most, only a minor pathway for the spread of voltage within the retinal microvasculature.

At present, it is uncertain whether the electrotonic architecture at non-retinal locations in the circulatory system is static or is dynamic. Since angiotensin's inhibition of axial transmission is a newly recognized action of this vasoactive molecule, it is not known whether this vasoactive signal modulates electrotonic transmission elsewhere. With extensive documentation that the efficacy of voltage spread is not uniform within the system, it is clearly risky to extrapolate the findings of our study to non-retinal tissues. In fact, it is imperative that the electrotonic architecture be characterized for each vascular bed.

Functional specialization within the retinal microvasculature

Characterization of the electrotonic architecture of the retinal microvasculature provided further support for the emerging concept that there is functional specialization within this operational unit. Specifically, our finding that radial transmission between an abluminal cell and the endothelium is significantly more efficient in the capillaries than in the arterioles supports our proposal (Matsushita et al. 2010) that the capillary network is particularly well adapted for generating and transmitting an electrophysiological response to a vasoactive signal. Consistent with this functional concept, the relative efficacy of radial transmission in the capillaries results in a substantial majority of the voltage generated in a pericyte being transmitted to the underlying endothelium, which then provides a highly effective pathway for proximal transmission. In addition to having the highest efficacy of radial transmission, capillaries are known to have physiological attributes that enhance their ability to generate a voltage change in response to vasoactive signals. For example, because functional K_ATP channels are chiefly located in the capillaries, this portion of the microvasculature generates almost all of the hyperpolarizing K_ATP current induced by adenosine (Ishizaki et al. 2009). In addition, because capillaries have a relatively small outward K_IR conductance (Matsushita & Puro, 2006) and thereby a higher membrane resistance, a change in ion channel activity induced by a vasoactive signal evokes a relatively large voltage change. On the other hand, although capillaries effectively generate and transmit voltages, their dearth of functional voltage-dependent calcium channels (VDCCs) prevents VDCC-dependent vasomotor responses from occurring in the capillary network (Ishizaki et al. 2009). Rather, the capillary's specialization for generating and transmitting voltages is complemented by the proximal microvasculature's abundance of VDCCs (Matsushita et al. 2010), which can convert a capillary-generated voltage into a change in abluminal cell calcium and, thereby, changes in abluminal cell contractility, lumen diameter and blood flow.

Effect of branch points on electrotonic transmission

This study also established that at some, but not all, branch points in the retinal microvasculature, there is significant voltage dissipation. We found that a spreading voltage dissipates by ∼45% as it passes proximally from a branch of the capillary network to a tertiary arteriole. Transmission through the branch point of tertiary and secondary arterioles also causes a similar attenuation of voltage. Thus, as deduced previously from the study of larger, i.e. 50–75 μm diameter, arterioles of the ileal submucosa (Segal & Neild, 1996), the effect of branch points must be considered in an analysis of how voltage flows through a microvascular network. On the other hand, our study indicated that not all classes of branch points within the retinal microvasculature had a detectable impact on the spread of voltage. Specifically, in our dual recording experiments, the efficacy of distal-to-proximal transmission was not significantly affected by intra-capillary or intra-arteriole branches. At present, an explanation for the differing effects of branch points within the retinal microvasculature awaits further study, although it appears likely that the relative diameters (Segal & Neild, 1996) and capacitative loads of the parent and daughter vessels may determine how much, if any, voltage dissipation occurs. Also awaiting determination is the functional consequence of the voltage dissipation that occurs at the capillary–tertiary arteriole junction and also at the junction of tertiary and secondary arterioles. Our working hypothesis is that a transmission efficacy of <1.0 at the site where sister branches converge enhances the ability of the proximal vessel to integrate inputs from these distal sites.

Angiotensin-induced inhibition of electrotonic transmission

An important operational concept established by this study is that angiotensin modulates the electrotonic architecture of the retinal microvasculature. Previously, we raised this possibility based on voltage-clamp recordings showing that a decrease in the membrane capacitance (C_m) is associated with exposure of retinal capillaries to angiotensin. However, because a change in C_m provides, at best, only a grossly qualitative indication of the extent of coupling within a population of cells (Zhao & Santos-Sacchi, 1998), does not allow determination of transmission efficacies, cannot distinguish effects on radial and/or axial transmission and is affected by membrane events unrelated to intercellular communication (Gillis, 1995), it was necessary to use a more specific and sensitive assay. To eliminate the ambiguity of a C_m assay, the present study quantified electrotonic transmission between a pair of current-clamped recording pipettes. We found that during exposure to angiotensin, the rate at which a voltage decays as it spreads along a capillary, tertiary arteriole or secondary arteriole is increased from ∼5%/100 μm to ∼50%/100 μm. Indicative that angiotensin selectively inhibits axial transmission, neither the efficacy of radial transmission between abluminal cells and the endothelium nor transmission at branch points between a capillary and a tertiary arteriole or between a tertiary and secondary arteriole was significantly affected by this vasoactive molecule.

Although the widespread effect of angiotensin on axial transmission in the retinal microvasculature is consistent with our finding that AT₁ angiotensin receptors are expressed throughout this microvascular complex, the mechanism by which angiotensin inhibits axial transmission is uncertain. However, our experiments indicate that this effect does not require a change in the membrane resistance since exposure to angiotensin caused a 10-fold decrease in the efficacy of axial transmission not only in the capillaries where angiotensin induces a non-specific cation conductance, but also in the arterioles where only small currents are activated by this vasoactive molecule.

What are the functional consequences of angiotensin's inhibition of axial transmission? An operational result is that the electrophysiological response of the capillaries to angiotensin remains highly localized. Specifically, our recordings from intact and transected microvascular complexes demonstrated that the depolarization caused by angiotensin's activation of non-specific cation channels in the capillary network remains confined to the capillaries. It seems probable that in the presence of angiotensin, voltage changes generated in the capillaries by other vasoactive molecules (Puro, 2007) would also be prevented from spreading proximally. In this way, input to a capillary may evoke a local, rather than a global, change in blood flow. Based on these experimentally derived considerations, we propose that the action of angiotensin to inhibit axial transmission allows this vasoactive signal to regulate the geographical extent of the microvasculature's response to voltage-changing inputs.

Isolated retinal microvascular complexes: advantages and caveats

This analysis of the electrotonic architecture of the retinal microvasculature was based on the study of freshly isolated microvascular complexes. With this experimental preparation, it was relatively straightforward to obtain dual perforated-patch recordings from abluminal cells located at well defined locations in a retinal microvascular complex. Also, the ability to transect an isolated microvascular complex at the capillary–tertiary arteriole junction and also at the tertiary arteriole–secondary arteriole junction permitted detection of currents generated within the capillaries, tertiary arterioles or secondary arterioles. Furthermore, use of isolated microvessels allowed us to assess the effect of angiotensin in the absence of confounding effects mediated via non-vascular cells. However, although recordings from endothelial cells would have helped in our assessment of electrotonic transmission, this proved impractical since the vascular endothelium is extensively covered by abluminal cells whose anatomy and function were disrupted during the process of sealing a pipette onto an endothelial cell. Also, the technique of internal perfusion of retinal microvessels with toxic chemicals to selectively kill the endothelial cells has yet to be perfected. In addition, since the isolated microvessels were not internally perfused, the effects of intralumnial pressure on electrotonic transmission were not evaluated in this study. In the future, as additional quantitative data concerning the retinal microvascular become available, a better understanding of the physiology of this operational unit can be gained by the formulation of a detailed computational model, as has been done for skeletal muscle resistance arteries (Diep et al. 2005). Also awaiting future study is the characterization of the specific connexins that constitute the homocellular and heterocellular gap junction pathways within the retinal microvasculature. Future studies should also assess how the electrotonic architecture of the microvasculature of the retina is affected by diabetes, which disrupts gap junction function in the retinal microvessels (Oku et al. 2001) and causes sight-threatening complications that involve, at least in part, the retina's renin–angiotensin system (Fletcher et al. 2010). Finally, it must be noted that the conclusions based on this study of isolated vessels require in vivo verification, although technical advances will be required in order to assess electrotonic transmission within the retinal microvasculature in vivo.

Conclusions

In summary, this study has shown that the electrotonic architecture of the retinal microvasculature is characterized by highly efficient axial transmission within the capillaries, tertiary arterioles and secondary arterioles. On other hand, radial abluminal cell-to-endothelium transmission, as well as transmission at branch points between a capillary and its tertiary arteriole and between a tertiary arteriole and its secondary arteriole, is significantly less efficient. This study has also established that angiotensin selectively and profoundly inhibits axial transmission within capillaries, tertiary arterioles and secondary arterioles. As a consequence, the angiotensin-induced depolarization generated in the capillaries remains localized to the capillary network and fails to spread to the calcium channel-rich proximal portions of the microvasculature. Of general physiological importance, angiotensin's inhibition of axial transmission establishes the operational concept that the electrotonic architecture of the retinal microvasculature is not static, but rather is dynamically modulated by vasoactive signals.

Glossary

Abbreviations

AT₁

angiotensin receptor type 1

C_m

membrane capacitance

K_ATP

ATP-sensitive potassium channel

K_IR

inwardly rectifying potassium channel

NSC

non-specific cation

VDCC

voltage-dependent calcium channel

Author contributions

All experiments were performed at the University of Michigan by T.Z. and D.W.; they also contributed to the design of experiments, the analysis of data and the critical review of drafts of the manuscript. G.X. contributed to project planning and manuscript review. D.G.P. led in the conception and design of the project, in the analysis of the data, and in the writing of the manuscript. All authors gave final approval of the version to be published.