This manuscript describes a straightforward protocol for the isolation of arterioles from the rat retina that can be used in electrophysiological, calcium imaging and pressure myography studies.
The retina is a highly metabolically active tissue that requires a substantial blood supply. The retinal circulation supports the inner retina, while the choroidal vessels supply the photoreceptors. Alterations in retinal perfusion contribute to numerous sight-threatening disorders, including diabetic retinopathy, glaucoma and retinal branch vein occlusions. Understanding the molecular mechanisms involved in the control of blood flow through the retina and how these are altered during ocular disease could lead to the identification of new targets for the treatment of these conditions. Retinal arterioles are the main resistance vessels of the retina, and consequently, play a key role in regulating retinal hemodynamics through changes in luminal diameter. In recent years, we have developed methods for isolating arterioles from the rat retina which are suitable for a wide range of applications including cell physiology studies. This preparation has already begun to yield new insights into how blood flow is controlled in the retina and has allowed us to identify some of the key changes that occur during ocular disease. In this article, we describe methods for the isolation of rat retinal arterioles and include protocols for their use in patch-clamp electrophysiology, calcium imaging and pressure myography studies. These vessels are also amenable for use in PCR-, western blotting- and immunohistochemistry-based studies.
Understanding how blood flow is controlled in the retina is an important goal, since abnormal blood flow has been implicated in the pathogenesis of a variety of sight-threatening retinal diseases1,2,3,4. The retinal circulation, which supplies the inner retinal neurons and glial cells, has an end artery arrangement, with all of the blood from the retinal arteries and arterioles passing through the capillaries to the retinal venules and finally veins5. Blood flow in the retina is regulated by the tone of the retinal arteries and arterioles as well as the contractile activity of the pericytes located on the walls of the retinal capillaries and post-capillary venules6,7,8. The control of retinal vascular tone is complex and is modulated by a range of inputs from the circulatory system and surrounding retinal tissue, including blood gases, circulating molecules and hormones, and vasoactive substances released from the retinal vascular endothelium and macroglia9,10,11. The retinal arterioles are the small arterial branches of the retina and are composed of a single layer of vascular smooth muscle cells and an inner lining of longitudinally arranged endothelial cells12,13,14. These vessels form the main site of vascular resistance within the retinal circulation and therefore play an important role in the local control of retinal blood flow. Retinal arterioles regulate capillary blood flow in the retina by dilating or constricting their luminal diameter, mediated by changes in vascular smooth muscle contractility10,15,16. Understanding the molecular mechanisms through which retinal arterioles regulate retinal perfusion therefore requires preparations where the arteriolar smooth muscle cells can be accessed and studied in conditions as close to physiological as possible.
Ex vivo preparations of isolated retinal blood vessels provide access to the vascular smooth muscle cells, whilst still retaining their functionality and interconnectivity with the underlying endothelium. Most studies to date using isolated vessels have focused on large bovine or porcine arterial vessels (60 – 150 µm). These can be mounted in commercially available wire or pressure myograph systems to enable pharmacological interrogation of vascular smooth muscle cell contractile mechanisms17,18. Such preparations have greatly contributed to our knowledge of retinal vascular physiology under normal conditions. Few studies have used retinal arterioles isolated from small laboratory animals as their smaller diameter (~ 8 – 45 µm) prevents their use in conventional myography systems19,20,21,22. An important advantage, however, of using vessels from small laboratory animals is the wide availability of genetically modified, transgenic and retinal disease models. Small laboratory animals are also more amenable for in vivo intervention studies.
Here we describe straightforward protocols for isolating and cannulating rat retinal arterioles for pressure myography experiments. Ca2+ imaging and electrophysiology protocols using these vessels are also detailed. These can provide further insights into the regulation of vascular smooth muscle contractility and blood flow in the retina.
All experiments described here were performed in accordance with guidelines contained within the UK Animals (Scientific Procedures) Act of 1986 and were approved by the Queen's University of Belfast Animal Welfare and Ethical Review Body.
1. Isolation of Retinal Arterioles
Note: The following protocol is used to isolate retinal arterioles. This method is optimized for rat retinal arterioles but can be used with mouse retinas. The yield of vessels, however, is lower when using mice.
2. Arteriolar Pressure Myography
NOTE: Arteriolar pressure myography is carried out as follows using equipment detailed in Figure 1A, B and the Table of Materials.
3. Ca2+ Imaging
Note: Isolated retinal arterioles are prepared for conventional (microfluorimetry) and confocal Ca2+ imaging as follows (using equipment detailed in Table of Materials).
4. Patch-Clamp Electrophysiology
Note: Whole-cell and single-channel recording is possible from individual arteriolar smooth muscle cells still embedded within their parent arterioles as follows (using equipment detailed in Table of Materials).
Figure 6A shows a schematic drawing of the rat retinal vascular tree. The diameter ranges of the first order (30 – 45 µm), second order (20 – 30 µm), and pre-capillary arterioles (8 – 20 µm) has been confirmed experimentally in our laboratory by confocal imaging of rat retinal whole-mount preparations immunohistochemically labelled for α-smooth muscle actin (Figure 6B). Upon dissociation of the retina, primary, secondary and pre-capillary arterioles can be identified based on their caliber and the arrangement of the vascular smooth muscle cells. The first and second order arterioles appear visually similar under bright-field microscopy, but can be distinguished on the basis of their size (Figure 6C). The pre-capillary arterioles are the smallest arterial vessels in the preparation and are easily recognizable due to their sparser arrangement of vascular smooth muscle fibers. The isolated arterioles can be clearly differentiated from capillaries and venules within the isolation. Capillaries are apparent as a meshwork of small caliber (4 – 10 µm in diameter) vessels, while venules are thin walled and lack smooth muscle cell coverage (Figure 6C). Primary, secondary and pre-capillary arterioles are suitable for pressure myography, Ca2+ imaging and patch-clamp studies.
Figure 7 shows a pressure myography experiment where a primary rat retinal arteriole has been cannulated and the intraluminal pressure raised to 40 mmHg. The arteriole is then maintained at this pressure to allow for the development of myogenic tone prior to the addition of 0Ca2+ Hanks' containing 10 µM wortmannin to obtain the passive diameter of the vessel. Figure 7A shows photomicrographs of the arteriole at various time points during the course of the experiment. A full time-course record showing the changes in vessel diameter over time have been plotted in Figure 7B using custom made edge-detection software27. Immediately upon pressurization the vessel dilates, which is then followed by an active myogenic constriction that reaches a steady-state level after 15 min. Addition of 0Ca2+ Hanks'/wortmannin solution dilates the vessel back to a level similar to that observed immediately following the initial pressure step. As described above, drugs may be applied to the vessels before or following pressurization to investigate the mechanisms underlying the development and maintenance of myogenic tone in these vessels. Using this approach, we have previously shown that stretch-activated Transient Receptor Potential Vanilloid 2 (TRPV2) channels and L- and T-type voltage-gated Ca2+ channels play a central role in facilitating myogenic tone development in first and second order rat retinal arterioles20,21,22. We have also reported that large-conductance Ca2+ activated potassium channels (BK channels)19,28 and Kv1-containing voltage-gated K+ channels act to oppose myogenic activity in these vessels, since addition of specific inhibitors for each of these channels triggers vasoconstriction (Figure 7C).
Figure 8 shows examples of conventional and confocal Ca2+ imaging in retinal arterioles prior to and following exposure to the vasoconstrictor peptide, endothelin-1 (Et-1). In conventional fura-2-based recordings (Figure 8A), Et-1 (10 nM) elicits a biphasic increase in [Ca2+]i in the retinal arteriolar smooth muscle layer, comprising of an initial transient component followed by a lower sustained component. We have previously characterized the transient and sustained components as being due to Ca2+ release from the endoplasmic reticulum and Ca2+ influx from the extracellular space, respectively29. Fluo-4-based confocal Ca2+ imaging provides a more detailed picture of the effects of Et-1 at the cellular level. In these experiments, it is evident that the smooth graded changes in global [Ca2+]i observed in our microfluorimetry studies result from Et-1 stimulating the activation of repetitive asynchronous [Ca2+]i oscillations in the neighbouring retinal arteriolar smooth muscle cells along the vessel wall (Figure 8B, C;30).
Figure 9 shows examples of single-channel and whole-cell patch clamp recordings from retinal arteriolar smooth muscle cells. To date, we have carried out both on-cell and inside-out single channel patch clamp recordings22,28. Figure 9A, B for example, show an on-cell patch clamp recording prior to and following membrane stretch (generated by applying negative pressure to the patch pipette). This particular patch contains two channels which are activated by mechanical stretch. We have previously demonstrated that these currents can be inhibited using TRPV2 channel pore-blocking antibodies22. The unitary conductance of the channels (249.71 pS) is also consistent with these currents being mediated by TRPV231.
Whole-cell voltage-activated currents can be evoked using voltage step protocols. Figure 9C shows a family of voltage-activated A-type K+ currents recorded using such an approach. These currents become evident when other large currents present in these cells (e.g., BK and Ca+-activated Cl– currents) are blocked using specific pharmacological agents32,33. Currents through non- or weakly voltage-dependent ion channels are typically examined using voltage ramp protocols. We have used such protocols to identify and characterize TRPV2 currents activated by hypotonic stretch22 and channel agonists (Figure 9D) in retinal arteriolar smooth muscle cells.
Figure 10 shows dual pressure myography and confocal Ca2+ imaging applied in a primary rat retinal arteriole. Pressurization triggers increased frequency of [Ca2+]i oscillations in the individual smooth muscle cells similar to those observed with Et-1. We have previously shown that these oscillations are triggered by the summation of Ca2+ sparks (localized Ca2+ release events) and contribute to the generation of myogenic tone20.
Figure 1. Experimental setup for pressure myography in rat retinal arterioles.A) Experimental equipment including inverted microscope, in-line heater for bath perfusate, 3-axis mechanical and micro-manipulators, manometer, cannula holder and air table. B) Schematic diagram showing the optimal arrangement of the cannulating pipette and vessel of interest in the recording chamber. This diagram also illustrates the basic set-up of the pressurization equipment. C) Schematic diagram shown the process of anchoring and moving the vessel using additional tungsten wire slips held in fine forceps. (i) Drop 1st slip to occlude vessel. (ii, iii) Use additional slips to nudge the occluding slip (direction indicated by the arrow) and accompanying vessel into the optimum orientation shown in (iv). D) Schematic diagram showing optimal positioning of occluding tungsten wire slip, helper pipette and cannula in relation to the vessel as arranged prior to cannulation. The outlet of the drug delivery system is positioned at a distance of ~ 500 µm from the vessel to reduce movement artifacts during drug application. Please click here to view a larger version of this figure.
Figure 2. Cannulation process for pressure myography.A) Photomicrographs showing a retinal arteriole anchored down in the recording chamber prior to (i), during (ii)-(v) and following cannulation (vi). Blue arrow indicates direction of movement of the cannula while the green arrow indicates the direction of movement of the helper pipette. B) Photomicrograph showing optimal region for recording of myogenic activity during pressurization as highlighted in red. Note, adjustment of light and focus to increase contrast of vessel walls enables better tracking of vessels edges for automated analysis. Scale bar indicates 50 µm. Please click here to view a larger version of this figure.
Figure 3. Drug delivery. A) Equipment used for drug delivery showing solution reservoirs connected to a multi-channel delivery manifold controlled by 3-way taps and attached to a 3-axis mechanical manipulator. B) Diagram showing the optimal vertical positioning of the drug delivery manifold in relation to the recording chamber. C) Photomicrograph of a vessel anchored down in the recording chamber showing the ideal positioning of the drug delivery outlet. Scale bar represents 100 µm. Please click here to view a larger version of this figure.
Figure 4. Enzymatic digestion of retinal arterioles for patch clamp recording. Light micrographs of a retinal arteriole before (A) and after (B) enzymatic digestion. Note the physical separation (indicated by arrows) of the smooth muscle cells (SMCs) from the underlying endothelial cells (ECs). Smooth muscle cells suitable for patch clamping are indicated with an asterisk. Scale bar represents 10 µm. Please click here to view a larger version of this figure.
Figure 5. Single channel and whole cell patch clamp recording. A) Illustration of the test currents observed during the 'seal test' protocol when: (i) the patch pipette enters the external bathing medium, (ii) the pipette tip comes into contact with the vascular smooth muscle cell plasma membrane and (iii) suction is applied to the back of pipette to enable gigaseal formation. Following pipette capacitance compensation using the patch clamp amplifier, single channel currents may now be recorded in the on-cell configuration or a patch of membrane may be excised by rapidly withdrawing the pipette to create an 'inside-out' patch. B) Current changes associated with the development of electrical access to the cell interior during application of the amphotericin B whole-cell perforated patch clamp technique (i): (ii) As amphotericin B partitions into the membrane, access resistance falls and capacitance transients elicited during hyperpolarizing voltage steps (from -40 to -60 mV) become larger; (iii) once access resistance (Ra) has fallen to < 15 MΩ, which usually takes 5 – 10 min following gigaseal formation, experimentation is possible. C) Prior to whole-cell recording, access resistance and cell capacitance must be compensated using the relevant dials on the patch clamp amplifier. Values of access resistance and cell capacitance provided by automated functions within patch-clamp software can be used to help guide this process: (i) shows the capacitance transient prior to compensation taken from the highlighted region in B(iii); (ii) shows the reduction in the capacitance transient normally observed when the access resistance and capacitance are compensated; (iii) series resistance compensation should then be corrected up to 75% and access resistance and capacitance compensations fine-tuned such that the resulting transient should be relatively equal in amplitude above and below the plateau level of current during the step. (iv) Care should be taken to ensure that the access resistance is not over compensated (indicated by the presence of 'ringing' of the transient), which can occur sometimes if access continues to improve during the course of the experiment. Please click here to view a larger version of this figure.
Figure 6. Identification of isolated rat retinal arterioles. A) Illustration showing the arrangement of the rat retinal microvascular tree. B) Confocal images of rat retinal arterioles and venules within flat mount preparations stained for αSMA (red; nuclei are stained blue; scale bars represent 50 µm). C) Light micrographs of isolated 1°, 2° and pre-capillary arterioles, a capillary network and venule (scale bars represent 10 µm). Please click here to view a larger version of this figure.
Figure 7. Arteriolar pressure myography recordings. A) Images of a cannulated retinal arteriole showing (i) resting diameter (approximately 35.6 µm), (ii) dilation upon pressurization, (iii) development of myogenic vasoconstriction and (iv) dilation due to the addition of 10 µM wortmannin in 0Ca2+ Hanks' solution. Scale bar represents 10 µm. B) Time course plot of the vessel diameter for the full experiment shown in A (sampled at 2.5 frames/s). C) Diameter trace from a different arteriole under steady-state myogenic tone (diameter 38.7 µm) showing the effects of the Kv1 channel inhibitor correolide (10 µM) and the BK channel inhibitor iberiotoxin (100 nM) (sampled at 0.5 frames/s). Please click here to view a larger version of this figure.
Figure 8. Effects of Et-1 on [Ca2+]i signaling activity in rat retinal arterioles. A) Conventional fura-2-based recording showing the effects of Et-1 (10nM) on global [Ca2+]i. Basal [Ca2+]i was 82 nM in this 1° arteriole. B) Fluo-4-based confocal xt images showing the effects of Et-1 on [Ca2+]i at the cellular level. C) Plot of the normalized fluorescence intensity (F/F0) measured in cells as marked in B. This figure has been modified from Tumelty et al.30 with permission. Please click here to view a larger version of this figure.
Figure 9. Single channel and whole-cell patch-clamp recordings from rat retinal arteriolar smooth muscle cells. A, B) On-cell single channel recordings from the same membrane patch prior to (A) and during (B) the application of negative pressure (-45 mmHg) to the patch pipette. Two channels are present in the patch (unitary current 12.81 pA; unitary conductance 249.71 pS) which show a substantial increase in activation under stretch conditions. Selected regions from the upper panels (i) are shown on a faster time base in the lower panels (ii). C) Family of A-type K+ channel currents (i) recorded in whole cell mode in the presence of inhibitors of BK and Ca2+-activated Cl– channel inhibitors, elicited in response to 20 mV voltage step increments between -80 mV to +80 mV (ii). D) Whole cell currents (i) elicited by voltage ramps between -80 mV and +80 mV before (black line) and during (red line) application of the TRPV2 channel agonist delta-9-tetrahydrocannabinol (Δ9-THC). The voltage ramp protocol used is shown in the lower panel (ii). Please click here to view a larger version of this figure.
Figure 10. Ca2+ confocal imaging prior to and following pressurization of a rat retinal arteriole. Representative confocal xt scans (i) of retinal arteriolar smooth muscle cells under (A) unpressurized and (B) pressurized conditions obtained from separate regions of the same vessel. (ii) Changes in normalized fluorescence (F/F0) recorded in individual cells a – e, as indicated in (i), plotted against time. Asterisks indicate individual subcellular Ca2+ sparks which increase in frequency after pressurization and summate to trigger cellular Ca2+ oscillations. Please click here to view a larger version of this figure.
Compound (mM) | Low Ca2+ Hanks' | 0Ca2+ Hanks' | Normal Hanks' | Isolation enzyme Protease | Isolation enzyme Collagenase | Isolation enzyme DNAse | Quench solution | Rmin solution | Rmax solution | Cell attached Extracellular solution | Cell attached pipette solution | Whole-cell extracellular solution | Whole-cell pipette solution |
Water purity (resistance) | ≥15 MΩ.cm | ≥15 MΩ.cm | ≥15 MΩ.cm | ≥15 MΩ.cm | ≥15 MΩ.cm | ≥15 MΩ.cm | ≥15 MΩ.cm | ≥15 MΩ.cm | ≥15 MΩ.cm | ≥15 MΩ.cm | ≥18 MΩ.cm | ≥15 MΩ.cm | ≥18 MΩ.cm |
NaCl | 140 | 140 | 140 | 140 | 140 | 140 | 140 | 140 | 140 | 140 | |||
kCl | 6 | 6 | 6 | 6 | 6 | 6 | 6 | 6 | 6 | 6 | 138 | ||
D-Glucose | 5 | 5 | 5 | 5 | 5 | 5 | 5 | 5 | 5 | 5 | 5 | 5 | |
MgCl2 | 1.3 | 1.3 | 1.3 | 1.3 | 1.3 | 1.3 | 1.3 | 1.3 | 1.3 | 1.3 | 1.3 | 1.3 | 1 |
HEPES | 10 | 10 | 10 | 10 | 10 | 10 | 10 | 10 | 10 | 10 | 10 | 10 | 10 |
CaCl2 | 0.1 | 2 | 0.1 | 0.1 | 0.1 | 10 | 2 | 2 | 2 | 0.2 | |||
EGTA | 4 | 0.5 | |||||||||||
MnCl | 10 | ||||||||||||
Ionomycin | 0.01 | ||||||||||||
KOH | 140 | 130 | |||||||||||
D-Gluconic acid | 130 | 130 | |||||||||||
NaOH | 10 | ||||||||||||
Penitrem A | 0.0001 | 0.0001 | 0.0001 | ||||||||||
4-aminopyridine | 10 | ||||||||||||
Nimodipine | 0.01 | 0.01 | |||||||||||
Fluoxetine | 0.1 | ||||||||||||
9-anthracenecarboxylic acid | 1 | ||||||||||||
Amphotericin B | 300-600 μg mL-1 | ||||||||||||
Protease Type XIV | ~0.01 % (0.4-0.6 mg/40 mL) | ||||||||||||
Collagenase Type 1A | 0.1 % (6 mg/60 mL) | ||||||||||||
DNAse I | 20 KU (20 μL of 1 MU/mL stock in 20 mL) | ||||||||||||
pH | 7.4 | 7.4 | 7.4 | 7.4 | 7.4 | 7.4 | 7.4 | 7.4 | 7.4 | 7.4 | 7.4 | 7.4 | 7.2 |
titrated with | NaOH | NaOH | NaOH | NaOH | NaOH | NaOH | NaOH | NaOH | NaOH | Tris | Tris | NaOH | KOH |
Table 1. Composition of solutions used in experimentation including examples of external and pipette solutions used for single channel (TRPV2) and whole cell (A-Type) currents.
The protocols described above require practice but should be achievable with minimal troubleshooting. On an average day we would obtain 6 – 8 usable arterioles from the isolation and achieve 3 – 4 successful experiments. If problems are encountered, however, there are some steps that can be taken to help improve success rates. Occasionally we have found, particularly when using younger rats (< 8 weeks), that the yield of arterioles can be low. To circumvent this problem, we would suggest centrifuging the retinal tissue (10 – 30 s at 500 x g) between each of the trituration steps in steps 1.12 – 1.14. This often helps to improve the yield of vessels but will also increase the amount of cell debris within the preparation.
When cannulating vessels for pressure myography studies it is important to check the arterioles carefully for any side-branches that may have been cleaved close to the bifurcation site. This represents a common cause of leakage and loss of pressure during experimentation. The main issue that can arise with the [Ca2+]i protocols is the level of dye loading. Poor dye loading leads to low signal-to-noise ratios, while overloading results in disruption of normal Ca2+ homeostasis. To reduce the likelihood of either of these issues, a small aliquot of retinal homogenate can be removed, and isolated vessels checked periodically during the loading protocol. When undertaking patch-clamp experiments, the success rate of gigaseal formation is highly dependent on how well the basal lamina has been digested. When initially testing this protocol or using new lots of enzymes it may be necessary to adjust the concentration/duration of enzyme digestion. Carefully monitoring the separation of the endothelial and smooth muscle layers will ensure sufficient digestion to remove enough basal laminal to gain access. Careful monitoring is also necessary to avoid over-digestion, which can manifest as the vessel begins to constrict. If this occurs, the smooth muscle cells are often too fragile for patch clamp recording. When applying enzymes, it is important to include DNAse I to ensure removal of strands of DNA liberated from damaged cells during the isolation process. DNA fragments are sticky and cause the forceps to adhere to the arteriole during the final stages of the cleaning process (step 4.4), often resulting in vessel loss. Cleaning of vessels is technically difficult and is best performed at high magnification (20X) with gentle sweeping motions of closed forceps of the finest possible tip diameter. Between sweeps, clean the forceps with lab roll.
As highlighted earlier, a key motivation behind the development of the protocols outlined in this manuscript was to better understand why blood flow is disrupted during retinal vascular diseases. Most of our work to date has focused on diabetic eye disease28,33. Arterioles can be isolated from the retinas of experimental rodent models of diabetes using the methods described in section 1. When experimenting on isolated retinal arterioles from diabetic animals, it is important to try to closely replicate the hyperglycemic conditions experienced by the vessels in vivo. For this reason, we would normally raise the D-glucose levels in our isolation and experimental solutions to 25 mM. Thickening of vascular basement membranes is a well-recognized phenomenon in retinal vessels during diabetes34,35. When using animals with prolonged diabetes (> 1 months disease duration), increased enzyme concentrations or digestion times are often needed to enable the application of patch-clamp recording methods.
An important limitation of using ex vivo isolated retinal arterioles to study retinal vascular physiology and pathophysiology is the loss of the surrounding retinal neuropile. Although removal of the retinal glial and neuronal cells enables easy access to the retinal vascular smooth muscle cells for cell physiology studies, the response of the vessels to vasoactive mediators can change dramatically in the absence and presence of retinal tissue. The actions of adenosine tri-phosphate (ATP), for example, provide a good illustration of this point. In isolated rat retinal arterioles, addition of ATP triggers a robust constriction of the vessels36, while in the presence of an intact neuropile, the vessels dilate37. Therefore, wherever possible, we usually try to validate key findings from our isolated arteriole preparations using ex vivo retinal whole-mounts and in vivo measurements of vessel diameter and blood flow16,37. Of note, new methods have recently emerged for studying small arterioles and capillaries in whole perfused porcine retinas ex vivo38,39. Such preparations are likely to greatly improve our understanding of how the retinal neuropile regulates retinal arteriolar and capillary tone and how changes in retinal haemodynamics modulate neuronal activity in the retina.
Although the procedures described in this paper are focused on the use of isolated rat retinal arterioles for understanding arteriolar smooth muscle cell physiology, we are currently developing protocols to also enable the study of endothelial cell function in these vessels. In preliminary work, we have been successful in modifying the enzymatic digestion of retinal arteriolar segments to yield viable endothelial cell tubes that are amenable to Ca2+ imaging and patchclamp recording studies. Cannulation of the isolated arterioles at both ends, enabling intraluminal delivery of drugs, could in the future also enable endothelium-dependent vasodilatory responses to be investigated in these vessels.
The authors have nothing to disclose.
Development of the protocols described in this paper was supported by grants from the following funding agencies: BBSRC (BB/I026359/1), Fight for Sight (1429 and 1822), The Juvenile Diabetes Research Foundation (2-2003-525), Wellcome Trust (074648/Z/04), British Heart Foundation (PG/11/94/29169), HSC R&D Division (STL/4748/13) and MRC (MC_PC_15026).
Beakers | Fisherbrand | 15409083 | Or any equilavent product |
Curved Scissors | Fisher Scientific | 50-109-3542 | Or any equilavent product |
Disposable plastic pipette/ transfer | Sarstedt | 86.1174 | 6mL is best but other sizes are acceptable; remove tip to widen aperture |
Dissecting microscope | Brunel Microscopes LTD | Or any equilavent product | |
Forceps | World Precision Instruments | Dumont #5 14095 | Any equivalent fine forceps |
Pasteur pipette | Fisherbrand | 11546963 | Fire polished to reduce friction but not enough to narrow the tip |
Petri dish | Sigma-Aldrich | P7741 | Or any equilavent 10cm product |
Pipette teat | Fisherbrand | 12426180 | Or any equilavent product |
Purified water supply | Merek | Milli-Q Integral Water Purification System | Or any equilavent system |
Round bottomed test tube | Fisherbrand | 14-958-10 B | 5 mL; any equilavent product |
Seratted forceps | Fisher Scientific | 17-467-230 | Or any equilavent product |
Single edge blades | Agar Scientific | T585 | Or any equilavent product |
Sylgard | Dow Corning | 184 | Or any pliable surface |
Testtube rack | Fisherbrand Derlin | 10257963 | Or any equilavent product |
Pressure myography | |||
3-axis mechanical manipulator | Scientifica | LBM-7 | x2 (or any equilavent product) |
3-way taps | Cole-Parmer | UY-30600-02 | Or any equilavent product |
Air table | Technical Manufacturing Corporation | Clean Bench | Or any equilavent product |
Analysis software | Image J | https://downloads.imagej.net/fiji/ | Free imaging software |
Analysis plugin | Myotraker | https://doi.org/10.1371/journal.pone.0091791.s002 | Custom plugin for imageJ Freely available for download |
Cannulation pipette glass | World Percision instruments | TW150F-4 | Or any equilavent product |
Computer | Dell | Optiplex 7010 | Or any equilavent product |
Digital Thermometer | RS | 206-3722 | Or any equilavent product |
Fluo-4-AM | Thermo Fisher Scientific | F14201 | Make 1 mM stock in DMSO |
Forceps | World Precision Instruments | Dumont #7 14097 | Any equivalent fine forceps |
Fura-2-AM | Thermo Fisher Scientific | F1221 | Make 1 mM stock in DMSO |
Glass bottomed recording chamber | Warner Instruments | 64-0759 | Or any equilavent product |
Helper pipette glass | World Percision instruments | IB150F-3 | Or any equilavent product |
In-line heater | Custom made | Commerical equilavent SH-27B Solution In-Line Heater from Harvard Apparatus | |
Inverted microscope | Nikon | Eclypse TE 300 | Or any equilavent product |
Manometer | Riester | LF1459 | Or any equilavent product |
Microelectrode puller | Sutter instruments | P97 | Or any equilavent product |
Microforge + Olympus CX 31 microscope | Glassworks | Fine Point F-550 | Or any equilavent product |
Micromanipulator | Sutter instruments | MP-285 | Or any equilavent product |
Multi-channel delivery manifold | Automate Scientific | Perfusion Pencil | Or any equilavent product |
Pipette filler | BD Plastipak | 1mL | Syringe heated and pulled to internal diameter of pipette |
Pipette holder | Molecular Devices | 1-HL-U | Or any equilavent product |
Suction pump | Interpet | AP1 | Converted aeration pump by reversing bellows |
Syringe | BD Plastipak | 20mL Luer-lok | Or any equilavent product |
Tungsten wire | Advent | W558818 | 75μm diameter |
Tygon Tubing | VWR | miscellaneous sizes | Or any equilavent product |
USB camera | Logitech | HD Pro Webcam C920 | Or any equilavent product |
Video capture software | Hypercam | version 2.28.01 | Or any equilavent product |
Water bath | Grant | Sub Aqua 12 Plus | Or any equilavent product |
x20 lens | Nikon | 20x/0.40 WD 3.8, ∞/0.17 | Or any equilavent product |
x4 lens | Nikon | 4x/0.10, WD 30, ∞/- | Or any equilavent product |
Y-connectors | World Percision Instruments | 14012 | Or any equilavent product |
Patch clamping | |||
3-way taps | Cole-Parmer | UY-30600-02 | Or any equilavent product |
3-axis mechanical manipulator | Scientifica | LBM-7 | Or any equilavent product |
Air table | Technical Manufacturing Corporation | Clean Bench | Or any equilavent product |
Amphotericin B | Sigma-Aldrich | A2411 | 3 mg disolved daily in 50 μL of DMSO (sonicate to dissolve) |
Amplifier | Molecular Devices | Axopatch 200a | Or any equilavent product |
Analogue digital converter | Axon | Digidata 1440A | Or any equilavent product |
Collagenase Type 1A | Sigma-Aldrich | C9891 | 0.1mg/mL in LCH |
Computer | Dell | Optiplex 7010 | Or any equilavent product |
Digital Thermometer | RS | 206-3722 | Or any equilavent product |
DNAse I | Millipore | 260913 | Working stock 1 MU mL-1 (10 MU diluted in 10 mL LCH) |
Faraday cage | Custom made | Any equilavent commerical product | |
Fine forceps | Dumont | No. 7 | Any superfine forcep |
Glass bottomed recording chamber | Warner Instruments | 64-0759 | Or any equilavent product |
Headstage | Molecular Devices | CV203BU | Or any equilavent product |
In-line heater | Custom made | Commerical equilavent SH-27B Solution In-Line Heater from Harvard Apparatus | |
Inverted microscope | Nikon | Eclypse TE 300 | Or any equilavent product |
Microelectrode puller | Sutter instruments | P97 | Or any equilavent product |
Microforge + Olympus CX 31 microscope | Glassworks | Fine Point F-550 | Or any equilavent product |
Micromanipulator | Sutter instruments | MP-285 | Or any equilavent product |
Multi-channel delivery manifold | Automate Scientific | Perfusion Pencil | Or any equilavent product |
Patching software | Molecular Devices | Pclamp v10.2 | Or any equilavent product |
Pipette filler | BD Plastipak | 1mL | Syringe heated and pulled to internal diameter of pipette |
Pipette holder | Molecular Devices | 1-HL-U | Or any equilavent product |
Protease Type XIV | Sigma-Aldrich | P5147 | 0.01mg/mL in LCH |
Single channel pipette glass | World Percision instruments | IB150F-3 | Or any equilavent product |
Suction pump | Interpet | AP1 | Converted aeration pump by reversing bellows |
Syringe | BD Plastipak | 20mL Luer-lok | Or any equilavent product |
Tungsten wire | Advent | W557418 | 50μm diameter |
Tygon Tubing | VWR | miscellaneous sizes | Any equilavent product to fit |
USB camera | Logitech | HD Pro Webcam C920 | Or any equilavent product |
Water bath | Grant | Sub Aqua 12 Plus | Or any equilavent product |
Whole cell pipette glass | Warner Instruments | GC150TF-7.5 | Or any equilavent product |
x20 lens | Nikon | 20x/0.40 WD 3.8, ∞/0.17 | Or any equilavent product |
x4 lens | Nikon | 4x/0.10, WD 30, ∞/- | Or any equilavent product |
x40 lens | Nikon | 40x/0.55, WD 2.1, ∞/1.2 | Or any equilavent product |
Y-connectors | World Percision Instruments | 14012 | Or any equilavent product |
Ca2+ imaging | |||
3-way taps | Cole-Parmer | UY-30600-02 | Or any equilavent product |
3-axis mechanical manipulator | Scientifica | LBM-7 | Or any equilavent product |
Acquisition software | Cairn Research Ltd. | Acquisition Engine V1.1.5 | Or any equilavent product; microfluorimetry |
Air table | Technical Manufacturing Corporation | Clean Bench | Or any equilavent product |
Analysis software for confocal imaging | Image J | https://downloads.imagej.net/fiji/ | Free imaging software |
Computer | Dell | Optiplex 7010 | Or any equilavent product |
Confocal microscope | Leica Geosystems | SP5 | Or any equilavent product; confocal |
Digital Thermometer | RS | 206-3722 | Or any equilavent product |
Fine forceps | Dumont | No. 7 | Any superfine forcep |
Glass bottomed recording chamber | Warner Instruments | 64-0759 | Or any equilavent product |
In-line heater | Custom made | Commerical equilavent SH-27B Solution In-Line Heater from Harvard Apparatus | |
Inverted microscope | Nikon | Eclipse TE2000 | Or any equilavent product; microfluorimetry |
Monochromator | Cairn Research Ltd. | Optoscan | Or any equilavent product; microfluorimetry |
Multi-channel delivery manifold | Automate Scientific | Perfusion Pencil | Or any equilavent product |
Pipette filler | BD Plastipak | 1mL | Syringe heated and pulled to internal diameter of pipette |
Software for confocal microscope | Leica Geosystems | LAS-AF version 3.3. | Or any equilavent product; confocal |
Suction pump | Interpet | AP1 | Converted aeration pump by reversing bellows |
Syringe | BD Plastipak | 20mL Luer-lok | Or any equilavent product |
Tungsten wire | Advent | W557418 | 50μm diameter |
Tygon Tubing | VWR | miscellaneous sizes | Any equilavent product to fit |
Water bath | Grant | Sub Aqua 12 Plus | Or any equilavent product |
x100 lens | Nikon | x100 N.A. 1.3 oil | Or any equilavent product; microfluorimetry |
x20 lens | Nikon | 20x/0.40 WD 3.8, ∞/0.17 | Or any equilavent product; microfluorimetry |
x20 lens | Leica Geosystems | HCX PL FLUOTAR, 20x/0.50, ∞/0.17/D | Or any equilavent product; confocal |
x4 lens | Nikon | 4x/0.10, WD 30, ∞/- | Or any equilavent product; microfluorimetry |
x4 lens | Leica Geosystems | C PLAN, 4x/0.10, ∞/-/✝ | Or any equilavent product; confocal |
x63 lens | Leica Geosystems | HCX PL APO, 63x/1.40 – 0.60 OIL CS ∞/0.17/E | Or any equilavent product; confocal |
Y-connectors | World Percision Instruments | 14012 | Or any equilavent product |
Pipettes | Specifications and settings for fabrication of pipettes for experimentation | ||
Helper pipette | World Percision instruments | IB150F-3 | Inner diameter: 0.86 mm Ramp: 261 Pressure: 300 Heat: 280 Pull : 0 Velocity: 56 Time: 250 No of cycles: 4 Tip diameter: 0.5-2 μm Polishing: yes Final Resistance: >10 MΩ |
Cannulation pipette | World Percision instruments | TW150F-4 | Inner diameter: 1.17 mm Ramp: 290 Pressure: 300 Heat: 280 Pull : 0 Velocity: 58 Time: 150 No of cycles: 4 Tip diameter: 3-10 μm Polishing: no Final Resistance: <1 MΩ |
On-cell patching | World Percision instruments | IB150F-3 | Inner diameter: 0.86 mm Ramp: 261 Pressure: 300 Heat: 280 Pull : 0 Velocity: 56 Time: 250 No of cycles: 4 Tip diameter: 0.5-2 μm Polishing: yes Final Resistance: >5 MΩ |
Whole-cell patching | Warner Instruments | GC150TF-7.5 | Inner diameter: 1.17 mm Ramp: 296 Pressure: 200 Heat: 287 Pull : 0 Velocity: 50 Time: 250 No of cycles: 4 Tip diameter: 2-3 μm Polishing: helpful but not necessary Final Resistance: 1-2 MΩ |