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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

This protocol presents two techniques to isolate the subcellular compartments of murine rod photoreceptors for protein analysis. The first method utilizes live retinae and cellulose filter paper to separate rod outer segments, while the second employs lyophilized retinae and adhesive tape to peel away rod inner and outer segment layers.

Abstract

Rod photoreceptors are highly polarized sensory neurons with distinct compartments. Mouse rods are long (~80 µm) and thin (~2 µm) and are laterally packed in the outermost layer of the retina, the photoreceptor layer, resulting in alignment of analogous subcellular compartments. Traditionally, tangential sectioning of the frozen flat-mounted retina has been used to study the movement and localization of proteins within different rod compartments. However, the high curvature of the rod-dominant mouse retina makes tangential sectioning challenging. Motivated by the study of protein transport between compartments, we developed two peeling methods that reliably isolate the rod outer segment (ROS) and other subcellular compartments for western blots. Our relatively quick and simple techniques deliver enriched and subcellular-specific fractions to quantitatively measure the distribution and redistribution of important photoreceptor proteins in normal rods. Moreover, these isolation techniques can also be easily adapted to isolate and quantitatively investigate the protein composition of other cellular layers within both healthy and degenerating retinae.

Introduction

Rod photoreceptor cells, tightly packed in the outermost layer of the neural retina, are an integral part of dim light vision. To function as faithful photon counters, rods utilize a G-protein-based signaling pathway, termed phototransduction, to generate rapid, amplified, and reproducible responses to single photon capture. This response to light ultimately triggers a change in the current at the plasma membrane and is subsequently signaled to the rest of the visual system1. As their name implies, each rod cell has a distinct rod-like shape and exhibits a highly polarized cellular morphology, consisting of an outer segment (OS), inner segment (IS), cell body (CB), and synaptic terminal (ST). Each subcellular compartment has specific protein machinery (membrane-bound and soluble), biomolecular features, and protein complexes that play crucial roles such as visual phototransduction, general housekeeping and protein synthesis, and synaptic transmission2,3.

Over 30 years ago, the light-dependent reciprocal movement of subcellular proteins, specifically transducin (away from the OS) and arrestin (towards the OS), was first observed4,5,6,7. Early on, this observed phenomenon was received with skepticism, due in part to immunohistochemistry's vulnerability to epitope masking8. In the early 2000s, stimulus-dependent protein translocation was confirmed by using a rigorous and arduous physical sectioning technique9. Serial tangential sectioning of the frozen flat-mounted rodent retinae followed by immunoblotting revealed that transducin9,10, arrestin11,12, and recoverin13 all undergo subcellular redistribution in response to light. It is believed that light-driven translocation of these key signaling proteins not only regulates the sensitivity of the phototransduction cascade9,14,15, but may also be neuroprotective against light damage16,17,18. Because light-driven protein transport in rods appears to be very significant to rod cell biology and physiology, techniques that permit the isolation of different subcellular compartments to determine protein distribution are valuable research tools.

Currently, there are a few methods aimed at isolating the rod subcellular compartments. However, these methods can be lengthy and difficult to reproduce, or require a sizable amount of retinal isolate. Rod outer segment (ROS) preparations via density gradient centrifugation19, for example, is commonly used to separate the ROS from retinal homogenate. This method is widely used for western blot, but the procedure is very time consuming and requires a minimum of 8-12 murine retinae20. On the other hand, serial tangential sectioning of frozen murine and rat retinae has been successfully implemented in isolating the OS, IS, CB, and ST9,11,13. However, this method is technically challenging due to the necessity of fully flattening the small and highly curved murine retina to align the retinal layers prior to tangential sectioning. Since there are a plethora of mouse models and transgenic mice recapitulating diseases of the visual system, the creation of a technique that reliably, quickly, and easily separates individual rod compartments holds promise in revealing the physiologic processes that occur in each specialized compartment and the mechanisms that underlie visual processes in health and disease.

To facilitate these investigations, we describe two peeling methods that isolate rod subcellular compartments more easily than current protocols. The first peeling method, adapted from a technique to expose fluorescently labeled bipolar cells for patch clamp recording21, employs cellulose filter paper to sequentially remove the ROS from a live, isolated murine retina (Figure 1). The second method, adapted from a procedure that isolates the three primary retinal cell layers from a chick22 and frog23 retina, utilizes adhesive tape to remove the ROS and rod inner segment (RIS) from a lyophilized retina (Figure 2). Both procedures can be completed in 1 h and are considerably user-friendly. We provide validation of the effectiveness of these two separation protocols for western blot by utilizing dark-adapted and light-exposed retinae from C57BL/6J mice to demonstrate light-induced translocation of rod transducin (GNAT1) and arrestin (ARR1). Moreover, using the tape peeling method, we provide additional evidence that our technique can be used to examine and address inconsistencies between protein localization data acquired by immunocytochemistry (ICC) and western blots. Specifically, our technique showed that: 1) the protein kinase C-alpha (PKCα) isoform is present not only in bipolar cells, but also in murine ROS and RIS, albeit in low concentrations24,25, and 2) rhodopsin kinase (GRK1) is present predominantly in the isolated OS sample. These data demonstrate the effectiveness of our two peeling techniques for separating and quantifying specific rod and retinal proteins.

Protocol

All experiments were performed according to the local institutional guidelines of the committee on research animal care from the University of Southern California (USC).

1. Live cell retinal peeling method

  1. Preparation of Ames' buffers, peeling papers, and dissection dish
    1. Using blunt tip iris scissors (or equivalent scissor type), cut the cellulose filter paper (Grade 413) into rectangles measuring approximately 5 mm x 2.5 mm. Store the cuttings for future use.
    2. Prepare 1 L of Ames' HEPES buffer by combining one bottle of Ames' Medium, 2.38 g HEPES, and 0.877 g of NaCl. Dissolve the reagents in a sterile glass bottle with distilled ultrapure water and adjust the osmolarity and pH to 280 mOsm and 7.4, respectively. Filter to sterilize (pore size 0.2 µm), seal the lid with paraffin film, and store at 4 °C.
    3. Prepare 1 L of Ames' bicarbonate buffer by combining one bottle of Ames' Medium and 1.9 g of NaHCO3. Dissolve the reagents in a sterile glass bottle with distilled ultrapure water and adjust the osmolarity and pH to 280 mOsm and 7.4, respectively. Filter to sterilize (pore size 0.2 µm), seal the lid with paraffin film, and store at 4 °C.
      NOTE: Ames' HEPES and Ames' bicarbonate buffers can be prepared in advance and stored for 1 week at 4 °C.
    4. At the bottom of a 35 x 10 mm Petri dish, create a lattice or checkerboard pattern with a scalpel (blade No. 11, 40 mm).
      NOTE: The textured bottom of the Petri dish holds the isolated, free-floating eye in place during the retinal dissection.
  2. Dissection of the retina
    NOTE: For this procedure, the buffer should be brought to and kept at room temperature (RT). Refresh the Ames' bicarbonate buffer every 15 min with fresh carbogenated Ames' bicarbonate buffer during the dissection. This maintains the necessary physiologic pH.
    1. Around 15-20 min before the dissection, oxygenate the Ames' HEPES buffer in a 100 mL laboratory or media bottle equipped with a tubing cap adapter and bubble the Ames' Bicarbonate buffer with 95% O2 and 5% CO2 in a tissue incubation chamber and in a 100 mL laboratory or media bottle equipped with a tubing cap adapter.
    2. Euthanize an equal number of both genders of C57BL/6J mice (2-3 months old) either by isoflurane inhalation or carbon dioxide (CO2) followed by cervical dislocation. Enucleate the eyes with curved scissors and place the eyes into the textured 35 x 10 mm Petri dish filled with bubbled Ames' bicarbonate buffer.
    3. Create an eye cup (Figure 1A) by puncturing a hole at the cornea-limbus junction with an 18 G needle. Cut along the perimeter of this junction using micro-dissection iris scissors to remove the cornea of each eye. Separate the lens and vitreous humor from each eye using tweezers and discard.
    4. Isolate the retina from the eye cup by carefully peeling the retinal pigmented epithelium (RPE)-sclera-choroid complex away from the neural retina. Discard the RPE-sclera-choroid complex. Ensure that the retina is not damaged during the dissection by handling only the edges.
    5. Transfer isolated retinae using a wide bore transfer pipette into the lid of a 60 x 15 mm dish filled with bubbled Ames' bicarbonate buffer.
    6. Orient the hemisphere-shaped retina concave up (the photoreceptors are facing down toward the bottom of the dish) and bisect each retina (through the optic nerve) using iris scissors or a scalpel (blade No. 10, 40 mm). Trim the curved edges of each half retina to make two rectangles (Figure 1A).
      NOTE: Minimizing the curvature of each halved retina allows for the retina to flatten better in subsequent peeling steps and produces an accurate peel of the rod outer segment (ROS) layer.
    7. Store the halved retinae in the tissue incubation chamber that is continuously bubbled with 95% O2 and 5% CO2.
      NOTE: Isolated retinae can be kept in the carbogenated Ames' bicarbonate buffer for ~24 h.
  3. Rod outer segment (ROS) collection by filter paper peeling
    NOTE: Refresh the RT oxygenated Ames' HEPES buffer every 15-20 min during the peeling process to maintain the necessary physiologic pH.
    1. Transfer one retinal rectangle from the tissue chamber with a wide bore transfer pipette and a 5 mm x 2.5 mm piece of filter paper into a 35 x 10 mm Petri dish filled with Ames' HEPES buffer bubbled with 100% O2.
    2. Orient the partitioned retina concave up and ensure the photoreceptors are facing down toward the bottom of the dish. Lightly grasp the sides of the halved retina with tweezers and move it on top of the filter paper. Once the retina is centered on the filter paper, move the filter paper upward so the halved retina touches the filter paper to create the ROS-to-filter paper adhesion (Figure 1B).
      NOTE: Make sure that the photoreceptor layer makes direct contact with the filter paper.
    3. Carefully lift the filter paper with the attached retina out of the Ames' HEPES buffer. Place the bottom side of the filter paper (the side without the retina) on a paper towel and dab 2-3 times to dry (Figure 1B).
    4. Add a drop of Ames' HEPES onto the side of the filter paper with the retina. Place the filter paper on the paper towel again to dry, as just described. Repeat this process two more times.
    5. Place the filter paper with the retina back into the Petri dish. Using tweezers, gently push all edges of the halved retina up and away from the filter paper on all sides.
    6. Delicately peel the retina away from the filter paper. Touch only the extreme perimeter of the retina during the peeling process to preserve the retina's structural integrity.
    7. Lift the filter paper from the Petri dish and remove excess liquid on a paper towel. Make sure the side that was in contact with the retina does not contact the paper towel.
    8. Place the filter paper with the partial ROS layer into a labeled tube on ice (Figure 1B). Keep the peeled retina submerged in Ames' HEPES buffer.
    9. Repeat the peeling process for this halved retina approximately 7-8 times to remove the entire ROS layer. After each peel, place the filter paper into the same tube, which will contain the isolated ROS from one halved retina.
      ​NOTE: Take care to not tear the halved retina when peeling it away from the filter paper, as the retina becomes thinner during this process.
    10. Place the tube containing all filter paper peels of the isolated ROS on ice for immediate use or freeze directly on dry ice and store at -80 °C.
    11. Use tweezers to transfer the leftover peeled retina (lacking ROS) into an appropriately labeled tube. Place the tube on ice for immediate use or freeze with dry ice and store at -80 °C.
    12. Repeat the peeling process for each halved retina and accurately label each tube.

2. Lyophilized retina peeling method

  1. Buffers, peeling medium, dissection dish preparation, and lyophilizer set-up
    1. In a 1 L storage bottle, prepare the Ringer's solution by adding 500 mL of distilled ultrapure water. Dissolve the reagents to reach a final concentration of 130 mM NaCl, 3.6 mM KCl, 2.4 mM MgCl2, 1.2 mM CaCl2, 10 mM HEPES, and 0.02 mM EDTA. Adjust the pH to 7.4 with NaOH, add ultrapure water to bring the final volume to 1 L, and adjust osmolarity to 313 mOsm.
    2. Prior to the use, filter the Ringer's solution with a sterile filter (pore size 0.2 µm), seal the lid with paraffin film, and store at 4 °C.
    3. Using blunt tip iris scissors (or equivalent scissor type), cut the cellulose filter paper into rectangles measuring approximately 5 mm x 2.5 mm. Use a pencil to write on one side of the filter paper to create a unique label.
    4. Create a lattice or checkerboard pattern in the bottom of a 35 x 10 mm Petri dish using a scalpel (blade No. 11, 40 mm).
    5. Turn on the lyophilizer according to the manufacturer's specifications.
    6. Acquire liquid nitrogen in a Dewar flask or similar insulating vacuum flask.
  2. Retinal dissection
    1. Complete the retinal dissection as described in section 1 of the protocol (Figure 1A). Use cold Ringer's solution instead of Ames' buffer.
    2. Post dissection, store the halved, rectangular retinae in a 35 x 10 mm Petri dish filled with cold Ringer's solution and place on ice.
  3. Retinal sample preparation for lyophilization
    1. Place a 5 x 2.5 mm piece of filter paper and transfer a halved retina into the texturized 35 x 10 mm Petri dish filled with cold Ringer's buffer. Use a wide bore transfer pipette to move each retina between dishes.
    2. Use tweezers to carefully flip the retina so it is oriented concave up (the photoreceptors are pointing upward) and move it so it rests on top of the filter paper (Figure 2A).
      NOTE: Make sure the photoreceptors do not contact the filter paper (the retinal ganglion cell layer should be in direct contact with the filter paper) and the unique label is visible. To easily and quickly identify the retinal isolate, it is recommended that the unique label should be located on the underside of the filter paper.
    3. Lift the filter paper by the edges upward so the halved piece of retina touches the filter paper and continue to lift the filter paper out of the Ringer's solution.
    4. Place the bottom side of the filter paper (the side without the retina on it) on a paper towel and carefully dab 2-3 times to remove excess liquid (Figure 2A).
    5. Pick up the filter paper with tweezers and add a drop of cold Ringer's onto the side of the filter paper with the retina. Place the filter paper on the paper towel again to remove excess liquid. Repeat the process two more times.
      NOTE: These alternating wetting and drying steps ensure that the tissue is firmly attached to the filter paper.
    6. Store each adhered retina/filter paper sample in a Petri dish filled with cold Ringer's until ready to freeze all samples. Repeat this process for all halved retinae.
    7. Obtain a clean and dry 35 x 10 mm Petri dish (bottom only) and a piece of aluminum foil cut into a 3.5 x 3.5 inch square.
    8. Lift each sample out of the cold Ringer's buffer with tweezers. Ensure that the tweezers contact only the filter paper edges, avoiding the halved retina.
    9. Place the bottom side of each filter paper (the side without the retina on it) on a paper towel and remove excess liquid.
    10. Add a drop of cold 1x PBS (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4, pH 7.4) onto the side of the filter paper where the retina is adhered. Dab the filter paper on the paper towel to remove excess liquid.
      NOTE: Make sure that the moisture from the filter paper is sufficiently removed before freezing.
    11. Place all filter paper pieces into the 35 x 10 mm Petri dish. Use a lint free tissue paper to wick away any excess liquid surrounding the filter paper.
      NOTE: Do not overcrowd the Petri dish with samples. If more than four murine eyes are used, consider using a larger Petri dish or multiple smaller Petri dishes to hold the retina/filter paper samples.
    12. Tightly wrap the entire dish with aluminum foil. Ensure that the edges of the aluminum foil are secured and smoothly pressed into the bottom of the Petri dish. Puncture a handful of 0.1-0.2 mm holes into the aluminum foil lid (Figure 2A).
    13. Using metal tongs, gradually lower the aluminum foil covered Petri dish into the liquid nitrogen. Keep the samples in the liquid nitrogen (<10 min) until ready to lyophilize.
  4. Lyophilization
    1. Place the aluminum foil covered Petri dish into a freeze-drying flask and attach it to the lyophilizer machine following the manufacturer's protocol. Lyophilize the retinae for 30 min.
    2. Detach the flask from the lyophilizer and shut off the machine according to the manufacturer.
    3. Remove the aluminum foil and either work with the freeze-dried samples the same day or store them in properly labeled 1 mL microcentrifuge tubes. Place these tubes in a container (such as a 50 mL conical tube) filled with an anhydrous desiccant and store the samples at -80 °C.
  5. Rod outer and inner segment collection by adhesive tape peeling.
    1. Cut strips of clear adhesive tape (1-2 cm) and store strips on the edge of the tape dispenser/lab bench/etc. for quick use.
    2. Move one retinal sample to the lid of a plastic 60 x 15 mm Petri dish for peeling. Ensure to only grab the outermost edges of the filter paper and avoid touching the retina.
    3. Using blunt tip iris scissors (or equivalent scissor type), cut a small rectangular piece of tape to the approximate size of the tissue (1.5 x 2.5 mm).
    4. Carefully lay a small piece of tape on top of the lyophilized retina (Figure 2B) and apply slight pressure with the tweezers to make sure it bonds with the photoreceptor layer (the orange/pink-tinted top layer).
    5. Slowly peel the tape away. Both the rod outer segment (ROS) and rod inner segment (RIS) will be adhered to the tape (Figure 2B).
    6. Ensure that there is a thin white film at the fractured surface. This is RIS.To separate the RIS, place another piece of tape and push the tape down on the fractured surface (Figure 2B).
      NOTE: It may take more than one piece of tape to remove the entire thin white layer.
    7. Place the piece of tape with only the orange/pink layer into a microcentrifuge tube labeled +ROS.
    8. Place the tape or pieces of tape with the thin white layer into a microcentrifuge tube labeled +RIS.
      NOTE: Separating the lyophilized retina with tape takes practice. The amount of pressure applied to the tape will affect how the lyophilized sample will fraction. If too much pressure is placed on the tape over the sample, the whole lyophilized retina will peel off the filter paper and will stick to the tape. If too little pressure is added to the tape, the orange/pink top layer (+ROS) will not stick to the tape.
    9. Collect the leftover retinal tissue (bereft of ROS and RIS layers, -OIS) located on the filter paper by peeling off the thick, white layer from the filter paper using tape. Place the isolate into a tube labeled -OIS.
    10. Repeat these steps for each halved retinal sample.
    11. Store the isolated layers from the lyophilized retinae either at room temperature if performing protein quantification, gel electrophoresis, and protein immunoblots that same day, or store in a container (such as 50 mL conical tube) filled with anhydrous desiccant at -80 °C for later use.

3. Western blot sample preparation for peeling isolations

  1. Prepare RIPA lysis buffer
    1. In a 100 mL storage bottle, add reagents to obtain a final concentration of 50 mM Tris-HCl pH 7.4, 150 mM NaCl, 1% Triton X-100, 1% Sodium deoxycholate, 0.1% SDS, and 1 mM EDTA. Add 100 mL of deionized ultrapure water and mix thoroughly.
      NOTE: RIPA lysis buffer can be stored at 2-8 °C for 2-3 weeks. For longer storage, aliquot and store in the -20 °C freezer.
    2. On the day of the experiment, add a protease inhibitor cocktail (0.1 M PMSF, 1:1000 Aprotinin, and 1:1000 Leupeptin) to the RIPA lysis buffer and keep on ice.
  2. Filter paper peeling lysate preparation for western blot
    1. Ensure that tubes containing the filter paper peels and leftover peeled retinae are maintained on ice (for the experiment on the same day) or removed from -80 °C freezer storage and placed on ice.
    2. Remove any excess liquid from the bottom of the microcentrifuge tubes containing the filter paper peels. Quickly spin in a mini centrifuge for 2-4 s and discard any remaining liquid.
    3. Pipette 45-55 µL of cold RIPA buffer into tubes containing the filter paper isolate.
    4. Homogenize each sample with a clean, autoclaved pestle mixer for 1 min. Make sure the filter paper stays in contact with the pestle mixer. Keep the filter paper on the side of each tube rather than at the bottom.
    5. With tweezers, move the filter papers to the side of each tube and spin in the mini centrifuge for 2-4 s. This will remove the RIPA buffer from the filter paper.
    6. Remove dried filter paper and repeat for all filter papers within each tube if necessary.
    7. Transfer the homogenate to a new tube and label it appropriately.
      NOTE: This is the most time-consuming step, and if not completed properly, much of the sample will end up being absorbed by the filter paper.
    8. Pipette 60-80 µL of cold RIPA buffer into the individual tubes with the leftover peeled retinae.
    9. Homogenize each sample with a clean, autoclaved pestle mixer for 1 min.
  3. Tape peeling Lysate preparation for western blot
    1. Place the RT lyophilized samples or the -80 °C samples on ice.
    2. Pipette 50-70 µL of cold RIPA buffer into each tube containing +ROS and +RIS tape peels.
    3. Pipette 60-80 µL of cold lysis buffer into the -OIS tubes containing the remaining lyophilized retina, with the ROS and RIS removed.
    4. Homogenize each sample with a clean pestle mixer for 1 min. Ensure that the tape in the individual tubes stays in contact with the pestle mixer and lysis buffer.
    5. With sterilized tweezers, move the tape to the side of the tubes and spin with a mini centrifuge for 2-4 s. This will remove the liquid from the tape. Remove the dried tape from the tubes.
      NOTE: At this point, you may combine two half retinal isolates from the same retina for either the 'filter paper peeling' or 'tape peeling' method to ensure you have enough volume to perform a bicinchoninic acid assay (BCA).

Results

The present strategies were developed to provide relatively rapid and simple methods to isolate and analyze proteins among specific rod subcellular compartments for western blot analysis. We applied two sequential peeling techniques (Figure 1 and Figure 2) followed by immunoblotting to demonstrate that these methods could reliably be used to detect the known distribution of rod transducin (GNAT1) and arrestin (ARR1) in both dark- and light-adapted animals. To va...

Discussion

Many retinal diseases affect rod photoreceptor cells, leading to rod death and, ultimately, complete vision loss37. A significant portion of the genetic and mechanistic origins of human retinal degeneration have been successfully recapitulated in numerous mouse models over the years. In that context, the ability to easily and selectively separate individual rod subcellular compartments from the small mouse retina would greatly enhance our understanding of the localized biochemical and molecular un...

Disclosures

The authors declare that they have no competing interests.

Acknowledgements

This work was supported by NIH Grant EY12155, EY027193, and EY027387 to JC. We are thankful to Dr. Spyridon Michalakis (Caltech, Pasadena, USA) and Natalie Chen (USC, Los Angeles, USA) for proofreading the manuscript. We would also like to thank Dr. Seth Ruffins (USC, Los Angeles, USA) and Dr. Janos Peti-Peterdi (USC, Los Angeles, USA) for providing the necessary equipment to collect the author provided footage. Material from: Kasey Rose et al, Separation of photoreceptor cell compartments in mouse retina for protein analysis, Molecular Neurodegeneration, published [2017], [Springer Nature].

Materials

NameCompanyCatalog NumberComments
100 mL laboratory or media bottle equipped with a tubing cap adapterN/AN/A
100% O2 tankN/AN/A
1000mL Bottle Top Filter, PES Filter Material, 0.22 μmGenesee Scientific25-235
4X SDS Sample BufferMillipore Sigma70607-3
50 mL Falcon tubeFisher Scientific14-432-22
95% O2 and 5% CO2 tankN/AN/A
Ames’ Medium with L-glutamine, without bicarbonateSigma-AldrichA1420
CaCl2 (99%, dihydrate)SigmaC-3881
Drierite (Anhydrous calcium sulfate, >98% CaSO4, >2% CoCl2)WA Hammond Drierite Co LTD21005
Falcon Easy-Grip Petri Dish (polystyrene, 35 x 10 mm)Falcon-Corning08-757-100A
Falcon Easy-Grip Tissue Culture Dish (60 x 15 mm)Falcon-Corning08-772F
Feather Scalpel (No. 10, 40 mm)VWR100499-578
Feather Scalpel (No. 11, 40 mm)VWT100499-580
KCl (99%)SigmaP-4504
Kimble Kontes pellet pestleSigmaz359971
Labconco Fast-Freeze FlasksLabconcoN/A
LN2 (liquid nitrogen) + Dewar flask or similar vacuum flaskN/AN/A
MgCl2SigmaM-9272
Milli-Q/de-ionized waterEMD MilliporeN/A
Na2HPO4 (powder)J.T. Baker4062-01
NaCl (crystal)EMD MilliporeSx0420-3
NaHCO3Amresco0865
OmniPur EDTAEMD4005
OmniPur HEPES, Free AcidEMD5320
Parafilm MSigma-AldrichP7793
Reynolds Wrap Aluminum FoilReynolds BrandsN/A
Scotch Magic Tape (12.7 mm x 32.9 m)Scotch-3MN/A
Sodium deoxycholateSigma-AldrichD67501
Spectrafuge mini centrifugeLabnet International, IncC1301
Tissue incubation chamber (purchased or custom made)N/AN/A
Tris-HClJ.T.Baker4103-02
Triton X-100Signma-AldrichT8787
VirTis Benchtop 2K Lyophilizer or equivalent machineSP ScientificN/A
VWR Grade 413 Filer Paper (diameter 5.5 cm, pore size 5 μm)VWR28310-015
Whatman Grade 1 Qualitative Filter Paper (diameter 9 cm, pore size 11 μm)Whatman/GE Healthcare1001-090
Wide bore transfer pipet, Global ScientificVWR76285-362

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