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How do Proteins migrate in MES vs. MOPS

How do Proteins migrate in MES vs. MOPS



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My gels look significantly different in MES (2-(N-morpholino)ethanesulfonic acid) and MOPS (3-(N-morpholino)propanesulfonic acid). That is to be expected. What I don't understand is why the simple presence of the counter-ion significantly changes the velocity of certain proteins.

Is this due to the different pH (MES is usually run at pH 8.0, MOPS is usually run at pH 7.0) or is the charge distribution on the counter-ion responsible for everything?


Well, to rationalize everyone's comments, I think @leonardo is right.

This is a denaturing SDA PAGE gel. The migration of the SDS Micelles which are negatively charged, depends upon the shielding of the solution around it. The difference in mobility is because the SDS micelles will experience a slightly different field at pH ~6.2 (MES) vs 7.2 (MOPS).

The thought that these have the same charge would be right at exactly the pH corresponding to the pKa. For these two ions the proportions will not be the same when running 0.8 pH units from their respective pKas. the ion concentration goes as +/- log ([BH]/[B-]) and the charge environment of the buffer should not be the same. I think that the higher pH for MOPS running will tend to create more negative charge in the solution from MOPS- but also OH- in solution, slowing the mobility of the micelles and favoring the resolution of the larger proteins with MOPS.

This is all given even if the buffering ion concentration is the same.

I'm sure the gory details are buried in the musty tomes of some physical biochemistry journals deep in the library, but this is how it sounds to me.


Bookshelf

NCBI Bookshelf. A service of the National Library of Medicine, National Institutes of Health.

Lodish H, Berk A, Zipursky SL, et al. Molecular Cell Biology. 4th edition. New York: W. H. Freeman 2000.

  • By agreement with the publisher, this book is accessible by the search feature, but cannot be browsed.


Making the Rate of Protein Migration Proportional to Molecular Weight

The movement of any charged species through an electric field is determined by its net charge, its molecular radius and the magnitude of the applied field. But the problem with natively folded proteins is that neither their net charge nor their molecular radius is molecular weight dependent. Instead, their net charge is determined by amino acid composition i.e. the sum of the positive and negative amino acids in the protein and molecular radius by the protein’s tertiary structure.

So in their native state, different proteins with the same molecular weight would migrate at different speeds in an electrical field depending on their charge and 3D shape.

To separate proteins in an electrical field based on their molecular weight only, we need to destroy the tertiary structure by reducing the protein to a linear molecule, and somehow mask the intrinsic net charge of the protein. That’s where SDS comes in.


Introduction

In expansion microscopy (ExM), preserved biological samples are embedded within a swellable hydrogel and physically expanded isotropically, leading to optically transparent samples which allow for nanoscale resolution and aberration-free microscopy imaging on conventional diffraction-limited microscopes (Chen, Tillberg, & Boyden, 2015 ). Recently, two new ExM versions have been developed, termed protein retention expansion microscopy (proExM Tillberg et al., 2016 ) and expansion fluorescence in situ hybridization (ExFISH Chen et al., 2016 ), where protein (proExM) and RNA (ExFISH) molecules are chemically anchored to the hydrogel and probed upon expansion. This unit describes the most common scenarios for proExM and ExFISH sample preparation, as well as general guidelines for sample handling and image acquisition.

In proExM, proteins are covalently anchored to a hydrogel matrix with a commercially available small molecule (Acryloyl-X SE, or AcX for short Tillberg et al., 2016 ) that binds to primary amine groups on proteins (Fig. 1E, left) and is then incorporated into the hydrogel polymer during the process of polymerization (also known as gelation of a hydrogel schematized in Fig. 1A to 1C). An enzymatic digestion using proteinase K (ProK) or a high-temperature treatment with detergent mechanically disrupts the embedded sample, allowing for isotropic expansion in water. One can anchor native proteins to the gel using AcX and apply fluorescent antibodies post-expansion (post-expansion staining, Fig. 1C). This version uses a mechanical homogenization method employing high temperature and detergent, without protease. One can alternatively apply AcX to fixed specimens expressing fluorescent proteins and/or labeled already with fluorescent antibodies, incorporating these directly into the gel (pre-expansion staining, Fig. 1A, or pre-expansion fluorescent protein expression, Fig. 1B). Fluorescent proteins and antibodies are sufficiently resistant to proteolysis that they largely survive the ProK application. Utilizing a lens with 300-nm diffraction-limited resolution, the resulting ∼4.5× linear expansion of the embedded samples (∼90× volumetric expansion) allows imaging of the sample with ∼300 nm/4.5∼60- to 70-nm effective resolution.

In ExFISH, RNAs are covalently anchored to the hydrogel matrix with a small molecule, which we call LabelX (which can be easily synthesized by reacting two commercially available molecules), that binds to guanine (in RNA and DNA, Fig. 1E, right) and also to the hydrogel polymer (Chen et al., 2016 ). The ExFISH pipeline is shown in Figure 1D: samples are treated with LabelX, the swellable hydrogel polymer is formed, ProK is applied, and then the specimen is washed with phosphate-buffered saline (PBS). Afterwards, RNA-FISH probes are applied to the sample in a hybridization buffer to label RNA molecules of interest. The sample is then expanded and imaged in a low-salt buffer. This nonzero salt concentration, which allows RNA-FISH probes to remain stably bound, results in a ∼3× linear expansion (∼27× volumetric expansion), or an effective resolution for a 300-nm diffraction-limited lens of ∼300 nm/3∼100 nm.

This unit is structured into the following sections. In Basic Protocols 1 and 2, we describe proExM protocols for cells and intact tissues (of <200 μm pre-expansion thickness) that have been immunostained before expansion (pre-expansion staining proExM), or have fluorescent proteins expressed before expansion, as illustrated in Figure 1A,B. These two basic protocols are written independently, so that readers can start with either of the two, with minimal reference to the other. The next section, Basic Protocol 3, describes steps that help with processing of thick tissues or tissue slices (e.g., 200 to 500 μm thick, or potentially thicker), which have been immunostained or had fluorescent proteins expressed before expansion (as illustrated in Fig. 1A, B). Support Protocol 1 describes the protocol for immunostaining after expansion (post-expansion staining proExM, as illustrated in Fig. 1C). We also briefly discuss key considerations when choosing antibodies and dyes for proExM in Support Protocol 2. In Basic Protocols 4 and 5, we describe ExFISH methodology for the cultured cells and intact tissues, respectively. In the last protocol, Basic Protocol 6, we describe different mounting strategies for preparing expanded ExM samples for imaging on inverted confocal microscopes and the Zeiss Z.1 light-sheet microscope. This section on imaging applies to all ExM-processed samples, including products of the proExM and ExFISH protocols here described.

It may be helpful to note that sample preparation and imaging with proExM and ExFISH only require chemicals and microscopes found in a typical biology lab or imaging facility, with the exception of anchoring reagents and sodium acrylate, which are available commercially at modest cost. Almost all the protocol steps can be carried out at room temperature on a wetlab bench (unless otherwise noted in the protocol). Basic equipment required includes a benchtop vortex mixer, a shaker, a 37°C incubator (and a 60°C incubator for ExFISH of cultured cells), a desiccator, a 4°C refrigerator, and a –20°C freezer. All the materials and supplies used in this unit are commercially available. Unless otherwise noted, in this unit “water” specifically means “double-distilled water.”

1: proExM for Cultured Cells

Materials

  • 1 M potassium hydroxide (KOH) in water
  • 30% (v/v) ethanol in water
  • 10 mg/ml sterile fibronectin in Phosphate-buffered saline (PBS Current Protocols, 2001 )
  • Cells of interest, e.g., human embryonic kidney (HEK) cells
  • Cell culture medium suitable for the cell type of choice (e.g., Dulbecco's Modified Eagle Medium (DMEM) for human embryonic kidney (HEK) cells)
  • 4% (w/v) paraformaldehyde (PFA) in PBS or, for stronger fixation, a mixture of 4% (w/v) PFA and 0.1% (w/v) glutaraldehyde in PBS
  • 100 mM glycine in PBS
  • 0.1% (w/v) Triton X-100 in PBS
  • Blocking buffer: 5% (v/v) normal donkey serum in 0.1% (w/v) Triton X-100 in PBS (store up to 1 month or longer at 4°C) the serum should be from the animal host of the secondary antibodies, so other types of sera should be used instead of normal donkey serum if the secondary antibodies are from other animals
  • Primary and secondary antibodies for immunostaining
  • AcX/DMSO stock solution (see recipe)
  • Gelling solution for proExM of cultured cells (see recipe)
  • Digestion buffer (see recipe)
  • Cell culture substrates (e.g., coverslips of sizes, shapes, and surface modifications suitable for the culture protocol of choice), such as 12-mm round coverslips or a removable-chamber cover glass e.g., Sigma, cat. no. GBL 112358
  • Chamber cover glass removal tool (Sigma, catalog no. GBL 103259)
  • 35-mm petri dishes
  • Microcentrifuge tubes
  • Shaker
  • 22 × 22 mm No. 1.5 coverslips (0.15 to 0.19 mm thickness)
  • 22 × 22 mm No. 2 coverslips (0.19 to 0.23 mm thickness)
  • Microscope slides
  • Ice bath or cold block chilled to 4°C
  • Parafilm
  • Razor blade
  • Forceps
  • Nunc 4-well rectangular plate (Thermo Fisher, cat. no. 267061)
  • Paintbrush
  • Diamond scribe for cutting (scribing) thin glass
  • Dissection microscope

Cell culture

Cells can be cultured according to protocols of your choice. However, if there is no strong preference, we recommended culturing cells on a substrate separable from the culture well (e.g., on coverslips) or on a removable-chamber cover glass (e.g., Sigma, cat. no. GBL 112358) for easy gelation and sample handling in later steps.

The cell culture protocol described below uses adherent cell lines (e.g., HEK or COS7) and has demonstrated consistent results. If removable-chamber cover glasses are used for cell culture, start with the fibronectin glass surface modification in step 2.

1. Treat 12-mm round coverslips with 1 M KOH in water for 30 min, rinse with water three times, and store in 30% (v/v) ethanol in water.

2. Air-dry the cleaned 12-mm round coverslips in a sterile environment (i.e., biosafety cabinet), place in a 35-mm petri dish, and subsequently treat with 10 mg/ml sterile fibronectin in PBS for 30 min at 37°C. Remove the fibronectin solution immediately before cell plating.

3. Cells are plated and cultured on the surface of either the 12-mm coverslips or the removable-chamber cover glass to a confluency of 50% to 80% in a humidified incubator set at 5% CO2 and 37°C.

Cell fixation

Cells can be fixed using protocols of your choice. For instance, 4% (w/v) paraformaldehyde (PFA) in PBS can be used. For stronger fixation that preserves more ultrastructure, you might try using a mixture of PFA and a small percentage of glutaraldehyde (e.g., 4% (w/v) PFA plus 0.1% (w/v) glutaraldehyde in PBS), although glutaraldehyde-containing fixatives have been seldom used in expansion microscopy studies to date. The 4% (w/v) PFA in PBS is typically prepared freshly prior to the fixation.

The fixation protocol described below has demonstrated consistent results. Note that all the washing steps should be performed quickly to avoid sample dehydration.

4. Replace the cell culture medium with 4% (w/v) PFA in PBS and incubate for 10 min at room temperature.

CAUTION: This should be done in a chemical hood, as PFA is considered a potential carcinogen.

5. Remove the PFA solution and wash the cells for 5 min with 100 mM glycine in PBS to quench the fixation.

6. Remove the glycine solution and wash the cells twice for 5 min each with PBS.

7. We recommend moving to the next steps of the protocol immediately.

If necessary, fixed cells can be stored in the dark at 4°C in PBS for up to several weeks before the next steps of the protocol.

Immunostaining/immunohistochemistry (optional)

For immunostaining after fixation, follow protocols of your choice you can essentially perform conventional histology. The protocol described below has demonstrated consistent results with fixed cells prepared by the protocols described earlier.

8. Permeabilize the fixed cells by applying 0.1% (w/v) Triton X-100 in PBS at room temperature for 15 min. To reduce nonspecific antibody binding, apply blocking buffer at room temperature for 15 min.

9. Incubate the cells with primary antibodies in the blocking buffer, at the appropriate concentration. The incubation can be done on a shaker at low speed for a period ranging from 1 hr to overnight, at room temperature or at 4°C, depending on the antibody specifications.

10. Remove the antibody solution and wash four times, each time for 5 min with blocking buffer.

11. Incubate the cells with secondary antibodies in blocking buffer on a shaker at low speed for 2 to 4 hr at room temperature or overnight at 4°C. If using antibodies conjugated with dyes, see Support Protocol 2 for a discussion of dye compatibility with pre-expansion staining proExM.

12. Remove the antibody solution and wash four times, each time for 5 min with blocking buffer.

13. We recommend moving to the next steps of the protocol immediately.

If necessary, the samples can be stored in the dark at 4°C in PBS for up to several weeks before the next steps of the protocol.

Gelation

14. Prepare Acryloyl-X SE (AcX)/DMSO stock solution, monomer solution (Stock X) TEMED stock solution, and ammonium persulfate (APS) stock solution as instructed in Reagents and Solutions.

15. Replace the sample buffer (e.g., the blocking buffer if immunostained following the protocols above) with 0.1 mg/ml Acryloyl-X SE (AcX) in PBS.

This AcX solution is prepared by diluting the 10 mg/ml AcX/DMSO stock solution 1:100 in PBS.

16. Leave the cells in the 0.1 mg/ml AcX solution for 2 to 3 hr at room temperature.

Previous protocols suggested >6 hr or overnight, but we routinely find 2 to 3 hr to be more than enough time with no shaking.

17. When cells are cultured on a petri dish, the gelation chamber can be constructed in the meantime, by placing two spacers on a slide, separated from each other by a distance large enough so that the sample cover glass can fit in between (Fig. 2A). Thin and flat objects, such as stacks of coverslips, can serve as the spacers. For example, a stack of two No. 1.5 coverslips has a height of about 0.30 to 0.38 mm by changing the number or types of the coverslips, the height of the spacers can be adjusted accordingly to match the cell culture substrate (i.e., the sample cover glass) height. The spacer height should be kept close to the cell culture substrate height to avoid leaving excessive gel on top of the sample, but the total thickness of the gel [i.e., (spacer height) – (cell culture substrate height)] should be at least 0.15 mm for easier gel handling in later steps. A droplet of water (e.g., a few µl) can be applied to the spacers to adhere them to the slide (and, if multiple spacers, to each other). If a removable-chamber cover glass is used for the cell culture during steps 2 to 3, the cover glass itself and the gasket can serve as the gelation chamber and the spacer, respectively. In this case, the upper structure of the removable-chamber cover glass can be removed via a removal tool (Sigma, cat. no. GBL 103259). For the gelation chamber lid, a thick and robust coverslip (e.g., a No. 2 coverslip) wrapped in Parafilm can be used, with the Parafilm preventing the gel from sticking to the coverslip when removing the lid in later steps. Make sure the Parafilm surface is flat, clean, and free of folds. We recommend finishing preparation of the gelation chamber and the lid before moving to the next step.

18. After incubating the cells in AcX solution, wash the cells twice, each time for 15 min, with PBS. Begin thawing the components for the gelling solution for proExM—the Stock X, APS, and TEMED stock solutions. Keep the solutions chilled in an ice bath or on a cold block (chilled to 4°C).

19. Create the gelling solution for proExM now by mixing Stock X, water, TEMED stock solution, and APS stock solution in a 47:1:1:1 ratio (gelling solution also see recipes in Reagents and Solutions), in this order (i.e., APS last) in a microcentrifuge tube, and vortex for a few seconds.

The amount of gelling solution should be scaled up or down according to the size of the gelation chamber. For instance, 200 µl of gelling solution is enough to fill a gelation chamber with a 22 × 22 × 0.38 mm void. The mixed gelling solution should be kept in an ice bath or on a cold block (chilled to 4°C). Avoid excessive warming of the gelling solution by, for example, holding the lid of the microcentrifuge tube rather than the body of the tube. Once the gelling solution is prepared, we recommend completing the next few steps (until placing the gelation chamber in a 37°C incubator in step 24) within 5 min to prevent premature gelation.

20. Immediately after the previous step, remove the PBS from the cells using a pipet and add a small amount of the mixed gelling solution to the cells on the cell culture substrate. If a removable-chamber cover glass is used, remove the PBS and add the gelling solution to fill in the chamber, then continue to step 22.

21. Use a pair of forceps to pick up and place the cell culture substrate (i.e., the coverslip on which cells are cultured) with the cultured cells between the spacers in the gelation chamber. Make sure the cells are facing upward. Slightly press the cell culture substrate down, to hold it in place. If the gelling solution flows away, reapply a small amount of gelling solution to the cells.

22. Add a droplet (∼20 µl) of gelling solution to one side of the Parafilm-covered lid, and flip it over so that the droplet hangs from the surface due to surface tension (Fig. 2A). Slowly lower the lid so that the droplet will fuse with the gelling solution on the sample coverslip (Fig. 2B). Once fused, continue lowering the lid so it rests on the spacers. Make sure no air pockets are introduced inside the solution during this process.

23. Using a pipet with a fine tip, keep adding additional gelling solution to both open sides of the chamber until the space between the lid, spacers, and slide is filled with gelling solution (Fig. 2C).

Capillary forces should cause the gelling solution to evenly fill the void.

24. Place the gelation chamber in an incubator at 37°C for 1 hr for polymerization. Be careful not to tilt or shake the chamber during the transfer and the gelation.

Digestion

25. Take out the gelation chamber from the incubator.

26. Slowly insert a razor blade from the side between the lid and the spacers and pry the lid off. Once the lid is separated from the gel, use the razor blade to remove the spacers (Fig. 2D). If a removable-chamber cover glass is used, remove the lid, then use forceps to peel off the black gasket without removing the gel from the cover glass.

27. Using a razor blade, trim the gel to a small volume around the sample. This will keep the gel compact and aid in finding cells in later steps. Use the razor blade to remove the cell culture substrate and the gel, together, from the glass slide. If a removable-chamber cover glass is used, trim the gel but do not remove the gel from the cover glass.

You may want to trim the gel to have an asymmetric shape so that its orientation can easily be gauged in later steps simply by examination of the shape [see our video at https://www.youtube.com/watch?v=OksNCAJwxVI (proExM for tissues: gelation demonstration)].

28. Prepare digestion buffer as described in Reagents and Solutions (final concentration of ProK: 8 U/ml).

29. Since the gel will expand by about 1.5× during digestion, choose a suitably large container (e.g., petri dish) for the digestion. Also, use a separate container for each sample to avoid sample mix-up. Place the cell culture substrate with the gel in the container and fill it with digestion buffer of volume at least 10-fold excess of that of the gel (Fig. 2E). Immerse the gel in digestion buffer for at least 2 to 3 hr until it naturally peels off from the sample coverslip (Fig. 2F). If a removable-chamber cover glass is used, use a diamond scribe to separate each sample and its corresponding piece of the cover glass, by carefully scribing the cover glass and breaking each piece of glass off bearing its piece of gel. Place each piece of cover glass with the gel in a container and immerse in the digestion buffer as described above. A Nunc 4-well rectangular plate, for example, is convenient to contain several pieces of cover glass for digestion. The detached cell substrate can be removed using forceps, to leave only the cell-containing gel in the digestion buffer.

30. Leave the gel immersed in the digestion buffer overnight at room temperature in the dark.

Storage and expansion

31. If not done previously, remove the now-detached cell substrate (e.g., coverslip) from the digestion container carefully using forceps without disturbing the sample gel. Remove the digestion buffer and add PBS, and store the gel at 4°C in the dark if storage for later imaging is desired.

Be cautious not to suck up the gel while pipetting. The gel scatters light slightly at this stage, so changing the incident angle of light might help to locate it inside the solution.

32. If the gel needs to be transferred to another container (e.g., a larger container for trimming, or the sample holder for imaging), you can use a paintbrush of suitable size to pick up and move the gel. To avoid gel dehydration, add some PBS to the container.

33. Before expanding the sample, we recommend trimming the gel to the minimum size necessary. You can use a dissection microscope or a low-magnification wide-field microscope to locate the region of interest. To minimize sample movement, most of the surrounding liquid can be temporarily removed during trimming. Under the microscope, use a razor blade to cut and remove gel outside of the region of interest. We advise trimming the gel into an asymmetric shape to easily deduce the orientation. Make sure to trim the gel down to a reasonable size for handling and imaging, as the expansion will introduce an additional 2 to 2.5× (to a total of about 4.5×) linear increase in size. Transfer the gel with the regions of interest to a container (e.g., a petri dish of suitable size for expansion, or the sample holder for imaging) that is large enough to contain the expanded gel.

34. Fill the container and immerse the gel in water and wait for 20 min. Exchange the water and wait for another 20 min. Redo the exchange one more time (20 min in fresh water, three times in total).

The sample should be fully expanded and optically cleared as in Figure 3 and ready for imaging (these times are shorter than in previous publications, but reflect numbers that work for routine use in our lab). For long-term imaging, e.g., when imaging a large volume of specimen, sample mounting is necessary after the expansion and before the imaging (see Basic Protocol 6). Sample mounting attaches the sample to the sample holder and thus prevents it from drifting, and also can help keep the specimen within the working distance of the objective lens. For a quick check of the expanded sample under the microscope (<5 min), there is typically no need for sample mounting. On an inverted microscope, for example, you can use a pipet to temporarily (<5 min) remove the water around the gel to prevent it from sliding away.

35. If you need to transfer the expanded gel to a new container, it needs to be handled with care, as it is somewhat delicate in the expanded state. First remove most of the surrounding water and then use a brush to move the gel onto a clean glass coverslip placed in the same container. Next, gently lift the coverslip (with the gel on top) out of the container with forceps. The gel can be transferred by slowly brushing it off the coverslip to the new container. This method can be also used to flip the gel by inverting the coverslip bearing the gel, in case the gel is facing in the wrong direction.

2: proExM for Intact Tissues

The basic protocol for intact tissues differs slightly from that for cultured cells. Due to the thickness of the samples, intact tissues require a longer time for AcX diffusion. Furthermore, the gelling solution is supplemented with a polymerization inhibitor to allow for longer diffusion of the gelling solution through the samples before the polymerization begins.

Materials

  • Phosphate-buffered saline (PBS Current Protocols, 2001 )
  • 4% (w/v) paraformaldehyde (PFA) in PBS or, for stronger fixation, a mixture of 4% (w/v) PFA and 0.1% (w/v) glutaraldehyde in PBS
  • Sinking solution (see recipe)
  • Dry ice
  • 2-methylbutane
  • Optimal cutting temperature compound (OCT), M-1 or another embedding matrix
  • 0.1% (w/v) Triton X-100 in PBS
  • Blocking buffer: 5% (v/v) normal donkey serum in 0.1% (w/v) Triton X-100 in PBS (store up to 1 month or longer at 4°C): the serum should be from the animal host of the secondary antibodies, so other types of sera should be used instead of normal donkey serum if the secondary antibodies are from other animals
  • Primary and secondary antibodies for immunostaining
  • AcX/DMSO stock solution (see recipe)
  • Gelling solution for expansion of intact tissues (see recipe)
  • Digestion buffer (see recipe)
  • Vibratome or cryostat microtome
  • 22 × 22 mm No. 1.5 coverslips (0.15 to 0.19 mm thickness)
  • 22 × 22 mm No. 2 coverslips (0.19 to 0.23 mm thickness)
  • Microscope slides
  • Parafilm
  • Microcentrifuge tubes
  • Paintbrush
  • Ice bath or cold block chilled to 4°C
  • Razor blade
  • Forceps
  • Petri dishes
  • Dissection microscope

Tissue preparation

In this basic proExM tissue protocol, the thickness of the tissue or the tissue slice is assumed to be <200 µm in its thinnest dimension. For tissue samples with thickness above 200 μm, see Basic Protocol 3.

Tissues can be fixed using protocols of your choice. For instance, immersion or perfusion fixation with 4% (w/v) paraformaldehyde (PFA) in PBS can be used. For stronger fixation that preserves more ultrastructure, you might try a mixture of PFA and a small percentage of glutaraldehyde (e.g., 4% (w/v) PFA plus 0.1% (w/v) glutaraldehyde in PBS), although glutaraldehyde-containing fixatives have been less used in expansion microscopy protocols to date. The 4% (w/v) PFA in PBS solution is typically prepared freshly prior to the fixation.

1. After fixation, if tissues are thicker than 200 µm, cut them into <200 µm slices using a vibratome. Alternatively, for slicing in a cryostat, the tissue needs first to be cryoprotected by immersing it in sinking solution until the tissue sinks to the bottom. The tissue is then taken out and frozen using dry ice and 2-methylbutane. After embedding it in OCT, M-1 or another embedding matrix, the sample is ready to be sliced using a cryotome.

After either slicing method, we recommend moving to the next steps of the protocol immediately.

If necessary, fixed tissues or tissue slices can be stored in the dark at 4°C in PBS for up to several weeks before the next steps of the protocol.

Immunostaining/immunohistochemistry (optional)

For immunostaining after fixation, follow protocols of your choice. The protocol described below has demonstrated consistent results with fixed brain tissues prepared by the protocols described earlier.

2. Permeabilize the fixed tissue by applying 0.1% (w/v) Triton X-100 in PBS at room temperature for 15 min, then in blocking buffer at room temperature for 6 hr.

3. Incubate the tissues with primary antibodies in the blocking buffer. The incubation can be done on a shaker at low speed overnight at room temperature or at 4°C.

4. Remove the antibody solution and wash four times, each time for 30 min with blocking buffer, to remove unbound primary antibodies.

5. Incubate the tissues with secondary antibodies in the blocking buffer on a shaker at low speed overnight at room temperature or at 4°C. If using antibodies conjugated with dyes, see Support Protocol 2 for a description of dye performance and compatibility with pre-expansion staining proExM.

6. Remove the antibody solution and wash four times, each time for 30 min with blocking buffer to remove unbound secondary antibodies.

7. We recommend moving to the gelation steps immediately.

If necessary, the samples can be stored in the dark at 4°C in PBS for up to several weeks before gelation.

Gelation

8. Prepare Acryloyl-X SE (AcX)/DMSO stock solution, monomer solution (Stock X), TEMED stock solution, ammonium persulfate (APS) stock, and 4-hydroxy-TEMPO (4HT) solution as instructed in Reagents and Solutions.

9. Replace the sample buffer (e.g., the blocking buffer if immunostained following the protocols above) with 0.1 mg/ml Acryloyl-X SE (AcX) in PBS.

The AcX solution is prepared by diluting the 10 mg/ml AcX/DMSO stock solution 1:100 in PBS.

10. Leave the tissues in AcX solution for >6 hr (overnight is fine) at room temperature with no shaking.

11. Meanwhile, to construct the gelation chamber, place two spacers on a slide, separated from each other (Fig. 4A). Thin and flat objects, such as stacks of coverslips, can serve as the spacers. For example, a stack of two No. 1.5 coverslips has a height ranging from 0.30 to 0.38 mm, and by changing the number or types of the coverslips, the height of the spacers can be adjusted to accommodate the tissue thickness. The spacer height should be kept close to the tissue thickness to avoid leaving excessive gel on top of the sample, but the total thickness of the gel (i.e., the spacer height) should be at least 0.15 mm for easier gel handling in later steps. A droplet of water (e.g., a few µl) can be applied to spacers to adhere them to the slide (and, if multiple spacers, to each other). For the gelation chamber lid, a thick and robust coverslip (e.g., a No. 2 coverslip) wrapped in Parafilm can be used, with the Parafilm preventing the gel from sticking to the coverslip when removing the lid in later steps. Make sure the Parafilm surface is flat, clean, and free of folds. We recommend finishing preparing the gelation chamber and the lid before moving to the next step.

12. After incubating the tissues in the AcX solution, wash the tissues twice, each time for 15 min, with PBS. Begin thawing the components for the gelling solution for proExM: Stock X, 4HT, APS and TEMED stock solutions. Keep the solutions chilled in an ice bath or on a cold block (chilled to 4°C).

13. Add Stock X, 4HT, TEMED and APS in a 47:1:1:1 ratio in this order (i.e., APS last) into a microcentrifuge tube (gelling solution also see recipe in Reagents and Solutions), and vortex for a few seconds. The amount of gelling solution should be scaled up or down accordingly to the size of the gelation chamber and the size of the tissue or tissue slice and should be at least 100-fold excess in volume. For instance, at least 400 µl of gelling solution is used to incubate a 100-μm single coronal slice of mouse brain, enough to fill a gelation chamber with a 22 × 22 × 0.38 mm void, which can fit a full coronal mouse brain slice. To prevent premature polymerization of the gelling solution, APS should be added to the solution last before vortexing. In addition, the mixed gelling solution should be kept in an ice bath or on a cold block (chilled to 4°C). Avoid excessive warming of the gelling solution by, for example, holding the lid of the microcentrifuge tube rather the body of the tube. Finally, we recommend completing the next steps (until placing the microcentrifuge tube with the gelling solution and the sample at 4°C in step 15) within 5 min to prevent premature gelation.

14. Immediately after the previous step, use a soft paintbrush to transfer and immerse the tissues in the gelling solution in the microcentrifuge tube.

15. Keep the microcentrifuge tube containing the sample and the gelling solution at 4°C in the dark for 30 min. After the 30 min incubation, we recommend completing the next few steps (until placing the gelation chamber in a 37°C incubator in step 19) within 5 min to prevent premature gelation.

16. Use the paintbrush to transfer and place the tissue slice between the spacers in the gelation chamber. Make sure the slice is lying flat without wrinkles or distortions. We recommend first adding a droplet of 20 µl of the gelling solution in between the spacers, transferring the tissue slice into the droplet, and using the paintbrush to slowly flatten the tissue within the liquid environment, gently pressing it onto the glass surface.

17. Add a droplet of 20 µl of the gelling solution to one side of the lid, and flip it over so that the droplet hangs from the surface due to surface tension (Fig. 4A). Slowly lower the lid so that the droplet will fuse with the gelling solution on the sample coverslip (Fig. 4B). Once fused, continue lowering the lid so it rests on the spacers. Make sure no air pockets are introduced inside the solution during this process.

18. Using a pipettor and a fine pipet tip, keep adding additional gelling solution to both open sides of the chamber until the space between the lid, spacers, and the slide is filled (Fig. 4C).

Capillary forces will cause the gelling solution to evenly fill the chamber.

19. Place the gelation chamber in an incubator at 37°C for 2 hr for polymerization. Be careful not to tilt or shake the chamber during the transfer and the gelation.

Digestion

20. Take out the gelation chamber from the incubator.

21. Slowly insert a razor blade from the side between the lid and the spacers and pry the lid open. Once the lid is separated from the gel, continue to use the razor blade to remove the spacers in a similar fashion (Fig. 4D).

22. Using a razor blade, trim off any excess gel.

Trimming the gel will keep it compact and aid in finding the regions of interest in later steps. You may want to trim the gel to have an asymmetric shape so that its orientation can easily be gauged in later steps simply by examination of the shape [see our video at https://www.youtube.com/watch?v=OksNCAJwxVI (proExM for tissues: gelation demonstration)].

23. Prepare digestion buffer as described in Reagents and Solutions (final concentration of ProK is 8 U/ml).

We recommend preparing enough digestion buffer to have at least a 10-fold excess in volume compared with that of the gel.

24. Use a paintbrush to wet the gel and its sides with the digestion buffer. Wait for 10 to 30 sec, until the digestion buffer soaks the gel border.

This will aid in accessing the gel bottom with the paintbrush, so that the gel can be more easily peeled from the glass substrate.

25. Peel off the gel by gently probing the space between the gel and the glass surface with a fine paintbrush. Since the gel will expand to about 1.5× during digestion, choose a suitably large container, such as a petri dish, for the digestion step. For small tissue and tissue slice samples, 1.5-ml microcentrifuge tubes can be used as the container. Use separate containers filled with digestion buffer for each sample. Leave gels immersed in digestion buffer for >8 hr (overnight is fine) at room temperature in the dark (Fig. 4E, F).

Storage and expansion

26. Remove the digestion buffer and add PBS, and store the gel at 4°C in the dark if storage for later imaging is desired. Be cautious not to suck up the gel while pipetting.

The embedded tissue is slightly scattering at this stage, so changing the incident angle of light might help to locate it inside the solution.

27. If the gel needs to be transferred to another container (e.g., a larger container for trimming or the sample holder for imaging), you can use a paintbrush of suitable size to pick up and move the gel. To avoid sample dehydration, add some PBS to the new container.

28. Before expanding the sample, we recommend trimming the gel to the minimum size necessary. You can use a dissection microscope or a low-magnification wide-field microscope to locate the region of interest. To minimize sample movement, most of the surrounding liquid can be temporarily removed during trimming. Under the microscope, use a razor blade to cut and remove gel outside of the region of interest. We advise trimming the gel into an asymmetric shape to easily deduce the orientation. Make sure to trim the gel to a reasonable size for handling and imaging, as the expansion will introduce an additional 2× to 2.5× (to a total of about 4.5×) linear increase in size. Transfer the gel with the regions of interest to a container (e.g., a petri dish of suitable size for expansion, or the sample holder for imaging) that is large enough to contain the expanded gel.

29. Fill the container, immerse the gel in water, and wait for 20 min. Replace the water with fresh water and wait for another 20 min. Redo the exchange one more time (20 min in fresh water for three changes in total). The sample should be fully expanded and optically cleared (see Fig. 3 for an expanded mouse brain hemi-slice) and ready for imaging (these times are shorter than in previous publications, but reflect numbers that work for routine use in our lab). An example of pre-expansion and post-expansion images of a brain slice containing Brainbow 3.0-expressing neurons is shown in Figure 5. For long-term imaging, e.g., when imaging a large volume of specimen, sample mounting is necessary after the expansion and before the imaging (see Basic Protocol 6). Sample mounting attaches the sample to the sample holder and thus prevents it from drifting, and can also help keep the specimen within the working distance of the objective lens. For a quick check of the expanded sample under the microscope (<5 min), there is typically no need for sample mounting. On an inverted microscope, for example, you can use a pipet to temporarily (<5 min) remove the water around the gel to prevent it from sliding away.

30. If you need to transfer the expanded gel to a new container, it needs to be handled with care, as it is somewhat delicate in the expanded state. First, remove most of the surrounding water and then use a brush to move the gel onto a clean glass coverslip placed in the same container. Next, gently lift the coverslip (with the gel on top) out of the container with forceps. The gel can be transferred by slowly brushing it off the coverslip to the new container. This method can be also used to flip the gel by inverting the coverslip bearing the gel, in case the gel is facing in the wrong direction.

3: proExM for Thick Intact Tissue Samples (200 to 500 μm)

For tissues and tissue slices of 200 to 500 μm thickness, the basic tissue protocol (Basic Protocol 2) needs to be modified to allow for deeper penetration of reagents. In addition, the digestion time needs to be prolonged. We have successfully used this protocol to expand samples of up to 500 μm thickness expanding samples thicker than 500 μm may be possible as well.

Additional Materials (also see Basic Protocol 2 )

  • 2-(N-morpholino)ethanesulfonic acid (MES)–buffered saline (MBS): 100 mM MES/150 mM NaCl in water, adjusted to pH 6

1. Tissue sample generation is outlined in steps 1 to 7 of Basic Protocol 2, and can be used for thicker tissue samples.

2. For the AcX step (Basic Protocol 2, step 9), a slightly acidic buffer should be used to suppress the reactivity of the NHS ester of the AcX molecule to allow for deeper penetration into the tissue. We typically use MES-buffered saline (MBS) for this slightly acidic buffer. Dilute the 10 mg/ml AcX/DMSO stock solution 1:100 in the MBS. Leave tissue in the AcX in MBS solution for up to 24 hr at room temperature, depending on the thickness of the sample. We typically allow more time for thicker tissues. For instance, we leave a 500-μm brain slice in AcX/MBS solution for 24 hr.

3. For the gelling solution preparation (Basic Protocol 2, step 13), increase the amount of 4HT by 50% (i.e., 47: 1.5 ratio between the Stock X and 4HT) for longer inhibition of polymerization during the incubation. For the incubation (Basic Protocol 2, step 15), submerge the sample in the gelling solution for 45 to 60 min (instead of 30 min) at 4°C before transferring it to the gelation chamber.

4. For the digestion (Basic Protocol 2, step 25), a longer and stronger digestion is necessary. The ProK digestion can be carried out for a longer time and/or at higher temperature (up to 50° to 60°C), depending on the biological composition of the tissue and how strongly it is crossed-linked after fixation. For a 250-μm thick mouse brain slice, for example, a 2-day digestion at room temperature has yielded adequate digestion of the sample. In this case, the gelled brain slice was digested in digestion buffer (with ProK) at room temperature for 1 day, then the old buffer was exchanged with fresh digestion buffer (with ProK) and digested for another day.

1: Post-Expansion Immunostaining for proExM

Immunostaining can be performed after expansion, as illustrated in Figure 1C. One can perform the mechanical homogenization step using ProK, then add antibodies afterward. However, only some epitopes, such as YFP, can survive the ProK treatment and can be stained post-expansion most epitopes do not.

To preserve most epitopes, a milder method, such as protein denaturing by autoclaving in an alkaline detergent-rich buffer (or, alternatively, treatment with a mild protease that cuts at only a small number of sites within proteins, like LysC), can be used for mechanical homogenization (Fig. 1C, also see Supplementary Figure 1 of Tillberg et al., 2016 ). We describe the protocol for autoclave-mediated disruption below. For examples using various antibodies, see Supplementary Figure 1 of Tillberg et al., 2016 .

The following protocol refers to Basic Protocol 2, and an example using Thy1-YFP mouse brain slices and post-expansion anti-GFP staining is included (Fig. 6).

Additional Materials (also see Basic Protocol 2 )

1. Infuse a mouse brain slice with AcX and embed in a hydrogel as described in Basic Protocol 2.

After gelation, the sample can be peeled off from the glass substrate and temporarily stored in 1M NaCl solution (e.g., for periods of minutes to hours) before the next step.

2. Soak the sample in the alkaline buffer for 15 min, then replace the solution with fresh alkaline buffer of the same composition. Place the sample, immersed in the buffer, in an autoclavable container (e.g., polypropylene bottle), leaving the lid slightly open to allow for pressure relief. Autoclave the samples in the alkaline buffer on liquid sterilization mode, at a temperature of 121°C, for 1 hr. During this and subsequent steps, the sample will expand visibly.

3. Let the sample cool down to room temperature and wash twice, each time for 15 min, with PBS.

4. Incubate the sample with primary antibodies in 0.1% (w/v) Triton X-100 plus 5% (v/v) normal donkey serum in PBS (blocking buffer) on a shaker at low speed overnight at room temperature or 4°C.

The serum in the blocking buffer should be from the animal host of the secondary antibodies, so other types of sera should be used instead of the normal donkey serum if the secondary antibodies are from other animals.

5. Remove the primary antibody solution and wash four times, each time for 30 min, with blocking buffer.

6. Incubate the sample with fluorescently labeled secondary antibody of choice in the blocking buffer on a shaker at low speed overnight at room temperature or 4°C.

7. Remove the secondary antibody solution and wash four times, each time for 30 min, with blocking buffer.

8. Replace the buffer with PBS for storage (for up to several weeks), if desired. Expand to the final factor of ∼4.5× with three 20-min washes in water, and then proceed with imaging (see Basic Protocol 6).

For more detail on how to perform specific steps above, e.g., regarding sample handling, refer to corresponding steps in Basic Protocol 2.

2: Fluorescent Proteins, Antibodies, and Dyes for proexm

For the variation of proExM illustrated in Figure 1C (post-expansion staining proExM), any dye conjugated to the secondary antibody should be compatible with the protocol, as the dye is only administered at the end of the process.

For the variations of proExM illustrated in Figure 1A,B (pre-expansion staining proExM), most fluorescent proteins are compatible with the previously described basic proExM protocols (see Fig. 1b of Tillberg et al., 2016 ). For example, β-barrel (e.g., GFP-like) fluorescent proteins are protease resistant and can survive ProK digestion quite well (e.g., >50% fluorescence retention for many fluorescent proteins), but non-β-barrel fluorescent proteins (e.g., infrared fluorescent proteins based on bacteriophytochromes) are degraded by the ProK step. For antibodies conjugated with fluorescent dyes, most dyes are compatible with the basic proExM protocols (e.g., >40% fluorescence retention) except for the cyanine-family dyes (Fig. 1c of Tillberg et al., 2016 ). Cyanine-family dyes, such as Cy3, Cy5, and Alexa Fluor 647, are degraded during the polymerization reactions. For far-red dyes, accordingly, we recommend Atto 647N or CF 633.

4: Exfish for Cultured Cells

ExFISH enables expansion microscopy of RNA by using a small-molecule linker, which we call LabelX, to covalently anchor endogenous RNA molecules within specimens to the swellable gel of ExM. LabelX is analogous to AcX in proExM, but enables RNA molecules to be interrogated after expansion using RNA FISH. For ExFISH-processed cultured cells, single-molecule FISH (smFISH) is used to stain RNA molecules post-expansion.

Materials

  • Phosphate-buffered saline (PBS Current Protocols, 2001 )
  • 10% neutral buffered formalin (NBF EMS, cat. no. 15742-10)
  • 70% (v/v) ethanol in nuclease-free water
  • MOPS buffer (see recipe)
  • LabelX (see recipe)
  • VA-044 stock solution (see recipe)
  • Nitrogen gas source
  • Digestion buffer (see recipe)
  • Nuclease-free water
  • Wash buffer for smFISH (WA-10 see recipe)
  • smFISH hybridization buffer (see recipe)
  • smFISH probes: we primarily use commercially available smFISH probes (e.g., Stellaris ® RNA-FISH probes from LGC Biosearch Technologies). Alternatively, it is possible to design custom smFISH probes conjugated to a fluorophore of one's choice (Raj & Tyagi, 2010 ).
  • Parafilm
  • Tupperware container (modified as described in the text below) as an airtight chamber
  • 16-well Culturewell removable-chamber cover glasses from Grace Bio Labs (Sigma, cat. no. GBL112358)
  • Chamber coverglass removal tool (Sigma, cat. no. GBL 103259)
  • Vacuum desiccator and vacuum source
  • Glass microscope slides, 1-mm-thick
  • Kapton Tape or Scotch Tape
  • 60°C incubator
  • Razor blades
  • Diamond scribe for cutting (scribing) thin glass
  • Forceps
  • Nunc 4-well rectangular plates (e.g., Thermo Fisher, catalog no. 267061)
  • Epoxy glue

Cell culture preparation

Cells can be grown as desired to suit experimental needs. However, for ease of gelation and imaging, as well as handling, we have found 16-well Culturewell removable-chamber cover glasses from Grace Bio Labs (Sigma, cat. no. GBL112358) to be convenient for pre-expansion fluorescence in situ hybridization (ExFISH) imaging, as well as subsequent gelation and digestion. In cases where single-molecule FISH (smFISH) is performed before gelation and expansion, we have found it convenient to plate cells in 8-well Nunc Lab-Tek chambered cover glasses (Thermo Fisher, catalog no.155411).

Fixation and permeabilization of cultured cells

1. Wash cells once with PBS warmed to 37°C to avoid any heat shock.

2. Add 10% neutral buffered formalin and fix cells for 10 min at room temperature (all steps are at room temperature unless otherwise indicated). Wash with PBS twice, each time for 2 min at room temperature.

3. Replace buffer with 70% (v/v) ethanol in nuclease-free water for permeabilization. Fixed cells can be stored in 70% (v/v) ethanol in nuclease-free water at 4°C for up to 2 weeks. For immediate use, fixed cells should be permeabilized with 70% (v/v) ethanol in nuclease-free water for 1 hr at room temperature.

LabelX treatment of cultured cells

4. Rehydrate cells permeabilized with 70% (v/v) ethanol in nuclease-free water by washing twice with PBS for 5 min per wash at room temperature. Pre-incubate fixed cultured cells with 20 mM MOPS buffer for 5 min.

5. Prepare LabelX as described in Reagents and Solutions, and then dilute in 20 mM MOPS buffer to the desired concentration.

We have observed nearly complete RNA retention when using LabelX at a final Label-IT amine concentration of 0.006 mg/ml. For smFISH experiments, we recommend using a final Label-IT amine concentration of 0.006 mg/ml (i.e., a 1:150 dilution from the LabelX stock solution described in the recipe in Reagents and Solutions).

6. Remove the preincubation MOPS buffer and add LabelX in MOPS buffer to the fixed cultured cells. For cells grown in the removable-chamber cover glass, use 80 μl of LabelX in MOPS buffer. For cells grown in Nunc Lab-Tek chambered cover glasses, use a volume of 120 μl. Incubate overnight at 37°C. Afterwards, wash twice with PBS at room temperature for 5 min each.

At this stage, the cells can be temporarily stored at 4°C in PBS for up to a week.

Gelation

The Grace removable-chamber cover glass has an upper structure that can be removed with a chamber cover glass removal tool (Sigma, cat. no. GBL 103259). After removal of the chamber, a 1-mm silicone spacer remains with the cover glass and can be used to cast the gel (protocol described below, as well as in Basic Protocol 2). If cells are grown on a coverslip, then a chamber can be made using coverslip spacers (see Basic Protocol 2). If cells are grown in Nunc Lab-Tek chambered cover glasses, we have found that the bottom coverslip along with its 1-mm gasket can be removed from the rest of the plate, and used to cast the gel.

The gelation of fixed cultured cells for ExFISH is different from that of other ExM variants because VA-044, instead of APS, is used as a polymerization initiator. We found that the use of APS/TEMED yields a dim auto-fluorescent background that is significant in the context of smFISH, where the signals themselves are extremely dim. VA-044 is preferable because its use results in minimal auto-fluorescence, which is ideal for smFISH imaging. However, VA-044 has a higher decomposition temperature, and leads to a slower polymerization process. As a result, inhibition due to ambient oxygen can affect gel formation. We have found that degassing followed by perfusion with nitrogen displaces oxygen dissolved in the gelling solution and facilitates gel formation. Therefore, gelation is carried out under nitrogen atmosphere in a humidified chamber, which provides a nitrogen-filled environment and minimizes evaporation.

We typically use a Tupperware container, but any container that provides an airtight seal can be used for the humidified chamber (Fig. 7). We create two holes in the lid of the Tupperware using a syringe needle. To make the chamber humidified, we add a small amount of water to the bottom of the Tupperware. We position an empty 24-well plastic bottom plate at the very bottom of the Tupperware to provide a raised platform onto which we place the cover glass with the cells and gelling solution the water level should be much lower than the height of the 24-well plate so that the sample being gelled is kept well above the water.

7. For the Grace removable-chamber cover glass: Use the chamber cover glass removal tool to remove the top wells, leaving behind the black silicone gasket. Leave 40 μl of PBS inside the well to keep the cells hydrated. For the Lab-Tek chambered cover glass, first glue the bottom of the cover glass to a 1-mm-thick microscope glass slide for mechanical support, using epoxy glue. After the glue has cured, use a razor blade to pry apart the cover glass from the top wells leaving behind the plastic gasket. Leave 100 μl PBS inside the well to keep the cells hydrated.

8. Prepare gelling solution for ExFISH as follows.

Dilute the VA-044 to a final concentration of 0.5% (w/v) in monomer solution (i.e., dilute the stock 1:50 in monomer solution). For example, to prepare 1 ml of gelling solution, add 20 μl of VA-044 solution and 40 μl of water to 940 μl of monomer solution.

The gelling solution should be prepared at 4°C, and used for the next step immediately.

9. In a cold block, or on ice, distribute the gelling solution for ExFISH into 200-μl aliquots in microcentrifuge tubes. Degas for 10 min in a vacuum desiccator.

10. Remove any remaining PBS from the wells on the cover glasses and add the degassed gelling solution for ExFISH. For the Grace removable-chamber cover glass, add 40 μl of gelling solution. Add 200 μl of gelling solution if using the Lab-Tek chambered cover glass. Place the cover glass in a vacuum desiccator and degas for another 10 min. In the meantime, prepare a glass slide covered in Parafilm as the lid (see Fig. 2 and associated text, above). Make sure the Parafilm surface is flat, clean, and free of folds. The Parafilm prevents the gel from sticking to the lid when removing the lid in later steps. Remove the cover glass from the desiccator and immediately place the Parafilm glass slide directly on top of the cover glass. Any excess gelling solution might spill out to the sides. Proceed to the following steps without delay.

11. Place the cover glass onto the plastic platform in the humidified chamber. Seal the chamber (Fig. 7A,B).

12. Remove the tape covering the holes. Connect the inlet of the nitrogen line to a syringe tip and insert the tip through one hole (Fig. 7C). Slowly, turn on the nitrogen flow regulator to flush nitrogen into the chamber. Increase the flow until you can feel the airflow coming out of the other hole. Flush the chamber with nitrogen for 10 min. When done, remove the inlet and immediately seal both holes with tape again (e.g., Kapton Tape or Scotch Tape).

13. Place the chamber in a 60°C incubator for 2 hr to initiate the gelation (Fig. 7D). Be careful not to tilt or shake the chamber during the transfer and the gelation.

Digestion

14. When gelation is finished, take the humidified chamber out of the 60°C incubator and remove the cover glass. Using a razor blade, gently pry open the Parafilm-covered lid from the top of the wells. For the Grace Biolabs removable-chamber cover glass, gently remove the black silicone gasket by peeling it off using your hands or a pair of forceps. For the Lab-Tek chambered cover glass, do not attempt to remove the plastic gasket. At this point, you will see that gels will have formed in the individual wells on the cover glass.

15. For digestion, place the cover glass in a Nunc 4-well rectangular plate. Prepare digestion buffer with proteinase K as described in Reagents and Solutions. Add 6 ml of digestion buffer to the well containing the cover glass (if using any other container, add digestion buffer at at least 10 times the gel volume to completely cover the cover glass and gels). Make sure the gel is entirely immersed for at least 2 to 3 hr until it naturally peels off from the sample cover glass (Fig. 2F). If a removable-chamber cover glass is used, use a diamond scribe to separate each sample and its corresponding piece of the cover glass, by scribing the cover glass and breaking each piece of glass off bearing its piece of gel. Note that the cell culture substrate is placed so that the gel is on top of the substrate and the cultured cells are at the bottom surface of the gel. Optionally, the detached cell substrate can be carefully removed using forceps, to just leave the cell-containing gel in the digestion buffer.

16. Leave the gel immersed in the digestion buffer overnight at room temperature in the dark.

Storage and expansion

17. After digestion, gels can be expanded (temporarily, see below) in a petri dish, or any container large enough to hold the gels. To fully expand, wash gels with excess volume nuclease-free water three times with at least 1 hr per wash. Also see Basic Protocol 1.

SmFISH staining of cultured cells after expansion

18. Since the gels formed in either of the above chambered cover glasses have a thickness of ∼1 mm before expansion, the thickness of the expanded gels needs to be cut down to facilitate efficient diffusion of FISH probes through the gel. To shave down fully expanded gels to a thickness of 1 mm, we use 1-mm-thick microscope glass slides as spacers as follows. Prepare a large plastic dish to use as a cutting board. We often use the plastic covers of Nunc 4-well rectangular plates, although any similarly sized flat plastic surface will work. Use a small amount of epoxy to glue two 1-mm-thick glass microscope slides on their flat slide onto the plastic surface. Position the two glass slides such that they are aligned along their longest dimension with a 2.5- to 3-cm space in between. Allow the epoxy to harden.

19. Position a fully expanded gel in between the two glass slides on the plastic cover. The expanded gel should fit in between the two glass slides. If not, trim the gel with a razor blade. Position the gel such that the side of the gel with cultured cells is facing the bottom.

20. Place a razor blade across the glass slides so that it bridges the slides. Carefully slide the razor blade along the glass slides and through the gel to shave off everything except the bottom 1 mm of the gel containing the cells.

21. Carefully collect the shaved gel at the bottom containing the fixed and expanded cells and move it to PBS. If needed, at this point, shaved gels can be stored in PBS at 4°C for up to 1 month.

22. Proceed to smFISH staining, which can be performed in any container, though we often use glass bottom 24-well plates for the convenience of imaging right after staining.

23. Prepare gels by incubating with wash buffer (WA-10) for 30 min at room temperature.

24. Prepare probes by diluting in smFISH hybridization buffer at the concentration indicated by the vendor for smFISH. We typically carry out the hybridization at a total probe concentration of 100 nM. Vortex to mix.

25. Remove the wash buffer from the gels. Add the hybridization buffer with probes onto the gels. Add enough volume to completely cover the gel to be stained (for 24-well plates, we recommend 300 µl). Incubate overnight (or for >6 hr) at 37°C.

26. Wash gels twice with excess volume (e.g., 500 μl for 24-well plates) of WA-10 at 37°C with 30 min per wash.

27. Wash once with excess volume of PBS at 37°C for 30 min.

28. Imaging can be performed in PBS or any other buffer of choice [expansion factor is determined by salt concentration: using regular PBS results in ∼2× expansion using 0.02× PBS (diluted in water) results in ∼3× expansion while still preserving hybridization].

An example with HeLa cells is shown in Figure 8. Refer to Basic Protocol 6 for how to image expanded gels in the most common microscope setups. While the quality of the acquired image will depend on the imaging setup, wide-field imaging is generally recommended over confocal imaging due to the photobleaching of smFISH stains observed during confocal imaging.

The protocol for performing ExFISH in tissues is different from the cultured cell protocol in a few key aspects. First, the tissue ExFISH protocol requires signal amplification after performing FISH, because conventional smFISH is too dim for confocal imaging and similar 3-D imaging techniques needed for tissues. As a result, we use the hybridization chain reaction (HCR) to amplify the signal from FISH probes. Second, the gelling procedure is more similar to that described in the proExM protocol for tissues (Basic Protocol 2) than to the procedure for cultured cells. Third, higher concentrations of LabelX are used, because better RNA retention yields are observed in tissues at higher concentrations. Finally, after Label-X treatment, it is possible to use AcX to retain fluorescent proteins.

Materials

  • Nuclease-free water
  • Phosphate-buffered saline (PBS Current Protocols, 2001 )
  • 4% (w/v) paraformaldehyde (PFA) in PBS
  • 70% (v/v) ethanol in nuclease-free water
  • MOPS buffer (see recipe)
  • LabelX (see recipe)
  • Wash buffer for HCR-FISH (WA-20 see recipe)
  • HCR-FISH hybridization buffer (see recipe)
  • HCR-FISH probe design: probes for HCR-FISH are designed using Stellaris Probe Designer software from LGC Biosearch Technologies. 22-bp binding sequences spaced 2 bp apart are designed using the software. We often aim for at least 20 sequences targeting each transcript of interest. To design HCR-FISH probes, HCR initiator sequences (Choi, Beck, & Pierce, 2014 ) are appended to each binding sequence via a 2-base spacer (AA). The initiator sequences can be appended to either the 5′ end or 3′ end of each binding sequence depending on the orientation of the initiator sequences I1 initiators are appended to the 5′ end while I2 initiators are appended to the 3′ end. The HCR-FISH probes generated by this procedure are 60 bp in final length.
  • Amplification buffer (see recipe)
  • HCR Hairpins (purchased from Molecular Instruments each hairpin comes at 3 µM concentration in a storage buffer)
  • 5× SSCT (see recipe) and 0.05 × SSCT (see recipe)
  • Vibratome
  • 24-well culture plates
  • Additional reagents and equipment for transcardial perfusion (Chen et al., 2015 ), vibratome cutting (Chen et al., 2015 ), and proExM for intact tissues (Basic Protocol 2)

Tissue fixation and slicing

Tissues can be fixed using protocols of your choice. Below is a perfusion fixation and slicing protocol for mouse brain slices that has demonstrated consistent results.

1. Transcardially perfuse the mouse brain (Chen et al., 2015 ) with ice-cold PBS (5 to 10 ml) followed by ice-cold 4% (w/v) paraformaldehyde in PBS (∼30 ml).

2. Post-fix the brain in 4% (w/v) paraformaldehyde in PBS overnight at 4°C.

3. Wash the brain with PBS for 30 min once, before cutting slices.

4. Cut brain slices on a vibratome (Chen et al., 2015 ) in PBS.

The thickness of slices should be less than 200 µm to allow for efficient staining with FISH probes after expansion.

5. Slices can be stored in PBS if they are used immediately (within 1 to 2 days), or stored in 70% (v/v) ethanol in nuclease-free water for longer times at 4°C.

LabelX treatment, gelation, and digestion of tissues and tissue slices

6. If tissues or tissue slices were stored in 70% (v/v) ethanol in nuclease-free water, rehydrate by washing twice with PBS for 15 min per wash at room temperature. Otherwise, proceed with the rest of the protocol.

7. Pre-incubate tissues or tissue slices with 20 mM MOPS buffer for 30 min.

8. Prepare LabelX by diluting in 20 mM MOPS buffer at a final Label-IT amine concentration of 0.1 mg/ml (i.e., a 1:10 dilution from the LabelX stock solution).

9. Remove the pre-incubation MOPS buffer and add LabelX in MOPS buffer to the slices. For convenience, tissues or tissue slices can be processed in 24-well or 48-well plates. Use enough volume to cover the slices: for example, in a 24-well plate, four 50-µm sections can be incubated together in 250 µl of solution.

10. Incubate tissues or tissue slices with LabelX in MOPS overnight at 37°C. Afterwards, wash twice with PBS for 15 min per round.

11. Optional: To retain fluorescent proteins in tissue slices from transgenic specimens, it is possible to do an additional incubation with AcX. Incubate slices with 0.05 mg/ml AcX in PBS at room temperature for at least 6 hr. Wash twice with PBS for 15 min per wash.

12. Proceed to gelation and digestion as described in Basic Protocol 2.

Unlike the ExFISH gelation procedure for cultured cells, here, gelation with APS/TEMED can be carried out as in the proExM protocol because signal amplification with HCR renders the dim auto-fluorescence that arises from APS/TEMED negligible.

13. After digestion, wash gels with PBS twice, each time for 30 min. Gels can be stored in the dark in PBS at 4°C after this step for up to 2 months.

Staining gelled slices with HCR-FISH

While staining can be performed in any chamber, we often use 24-well plates.

14. Prepare gels by incubating with wash buffer (WA-20) for 30 min at room temperature.

15. Prepare probes by diluting in HCR-FISH hybridization buffer at the desired concentration (∼1 nM per probe). Vortex to mix.

16. Remove the wash buffer from the gels. Add the hybridization buffer with probes onto the gels. Add enough volume to completely cover the gel to be stained (for 24-well plates, 300 µl per well). Incubate overnight at 37°C.

17. Wash gels twice with excess volume (e.g., 500 µl for 24-well plates) of WA-20 at 37°C for 30 min per wash. For slices thicker than 100 µm before gelation, increase wash times to 45 min per wash.

18. Wash once with excess volume PBS at 37°C for 2 hr.

19. Wash once with excess volume of PBS at room temperature for 2 hr. For slices thicker than 100 µm before gelation, increase wash time to 4 hr. If needed, after this step, gels stained with FISH probes can be stored overnight in PBS at 4°C before HCR amplification.

HCR amplification (adapted from Choi et al., 2014 )

20. Pre-incubate slices with amplification buffer for 30 min at room temperature.

21. Prepare fluorescently labeled hairpins by snap cooling each hairpin (heat to 95°C for 90 sec, then cool to room temperature in a drawer for 30 min). For each 500 µl of total amplification buffer to be used (in the next step, we use 150 µl amplification buffer per well of a 48-well plate or 250 µl per well of a 24 well plate), 10 µl of each hairpin is needed.

22. Prepare hairpin solution by combining all snap-cooled hairpins in amplification buffer at room temperature, using the right volume of amplification buffer to get to the desired final volume. For example, to prepare 500 µl of total amplification buffer with two hairpins, add 10 µl of each hairpin to 480 µl of amplification buffer. Vortex to mix.

23. Remove the pre-amplification solution, add the hairpin solution to the gels, and incubate 3 to 6 hr at room temperature.

24. Stop amplification with four 30-min washes with 5× SSCT.

Expansion and imaging

25. At this point the samples can be imaged. To expand the samples, wash three times, each time for 10 min with 0.05× SSCT. The expansion factor can be tuned by altering the salt concentration imaging can also be performed in regular PBS for ∼2× expansion. An example mouse brain slice is shown in Figure 9. Refer to Basic Protocol 6 for how to image expanded gels in the most common microscope setups.

6: Imaging Expanded Samples

Materials

  • Expanded sample (see protocols above)
  • Ethanol
  • FISH probes
  • 0.1% (w/v) poly- L -lysine in water
  • Nitrogen source
  • Low-melting-point agarose
  • Superglue
  • Dissecting or wide-field microscope
  • Razor blade
  • Coverslips (e.g., 24 × 60 mm No. 1.5 (thickness: 0.15 to 0.19 mm))
  • Glass-bottom dishes or multi-well plates (e.g., Mattek, catalog no. P35G-0.170-14-C)
  • Paintbrush
  • Paper wipes (e.g., Kimwipes)
  • Inverted, upright, or light-sheet microscope

Sample expansion for imaging

1. Place the gel-embedded, digested sample in PBS (e.g., step 30 in Basic Protocol 1, end of step 25 in Basic Protocol 2, step 17 in Basic Protocol 4, or step 13 in Basic Protocol 5) in a petri dish.

2. Use a dissection microscope or a wide-field microscope with a low-magnification (e.g., 4× or 10×) objective to find the region of interest.

For ExFISH samples, this should be done before staining with FISH probes (i.e., before step 18 in Basic Protocol 4, or before step 14 in Basic Protocol 5).

3. Under the microscope, use a razor blade to remove excess gel (e.g., away from the region of interest).

Cultured cell ExFISH samples may also need to be trimmed axially to reduce thickness (see steps 18 to 21 in Basic Protocol 4).

4. Fully expand the proExM samples with water. ExFISH samples should be stained with FISH probes and expanded in diluted PBS for cultured cells, or diluted SSC buffer for tissues, following the protocols described earlier (i.e., steps 22 to 28 in Basic Protocol 4, or steps 14 to 25 in Basic Protocol 5).

Sample mounting

Mounting an expanded ExM sample on a stable surface can help prevent the sample from drifting during imaging, which is important for obtaining high-quality images. The best mounting method will depend on the geometries of your microscope and the sample holder (coverslip, Petri dish, or well of a multiwell plate, for example), as well as your imaging requirements such as imaging time and the objective type and magnification.

For a quick inspection (<5 min) of the expanded samples using an inverted or upright microscope with a dry objective, there is typically no need for sample mounting. You can use a pipet to temporarily (<5 min) remove the liquid around the gel to prevent it from floating or sliding away.

For long-term (>5 min) imaging or imaging that requires minimal sample drift (e.g., obtaining a z-stack), you might want to mount the sample by physically attaching the expanded samples to the sample holder. In the following sections, we describe three common mounting methods: agarose, poly- L -lysine, and superglue. For inverted microscopes, you can use poly- L -lysine to mount the gel to a stable surface. For upright microscopes, you can use agarose, poly- L -lysine, or superglue. For light-sheet microscopes, you can use superglue or poly- L -lysine.

The advantage of the poly- L -lysine method is that it provides a transparent interface between the gel and the surface of the sample holder. Therefore, it is suitable for microscope configurations that require imaging through the sample holder, such as an inverted microscope. In comparison, agarose can introduce optical scattering and aberration, and superglue is not transparent after curing. The advantage of the agarose method is that you can retrieve your sample after imaging, as the sample mounting is reversible. The sample can be shrunk (and then separated from the agarose) by placing in high-salt buffer after imaging. The advantage of the superglue method is its strong adhesion compared with the other methods. It is suitable for long-term imaging on an upright microscope or a light-sheet microscope, even for days.

Sample mounting with poly- L -lysine

5a. Coverslips, glass-bottom dishes, and glass-bottom multi-well plates can be used as sample holders. Slides or plastic multi-well plates can also be used, but the thickness of the sample holder may limit the choice of objectives and/or cause aberrations on inverted microscopes.

6a. Clean the surface of the sample holder by rinsing with water and then ethanol. After rinsing, let the holder dry in air.

7a. Soak the glass surface with 0.1% (w/v) poly- L -lysine in water for 20 min.

8a. Remove the 0.1% (w/v) poly- L -lysine solution and rinse the coated glass surface with water three times.

9a. Air-dry the glass surface for 1 hr in a clean environment or use nitrogen gas to blow-dry the surface.

  1. First, remove most of the liquid around the gel. Then place a clean coverslip next to the gel and use a paintbrush to slide the gel onto the coverslip.
  2. Next, lift the coverslip (with the gel on top) out of its container with a pair of forceps.
  3. Before transferring the gel to the poly- L -lysine-modified surface, the gel surface that is to be attached needs to be dried for good adhesion. You can use a small piece of a Kimwipe to wick away excess liquid. Start absorbing excess liquid from the sides of the gel and then carefully use, for example, the tip of the paper wipe to absorb the liquid in between the bottom of the gel and the coverslip.

Use the paintbrush to slide the gel off of the coverslip onto the poly- L -lysine-modified surface. Be aware of the orientation of the gel in the context of your later microscopy work: for example, the side facing toward the sample holder will be more easily reached by the objective (e.g., within the objective working distance) on an inverted setup. In contrast, on an upright microscopy setup, the side facing away from the sample holder will be more accessible to the objective. Therefore, if the region of interest is closer to one particular side of the gel, place that side closer to the objective when mounting the sample. This practice becomes important for expanded samples, as the sample thickness and the gel thickness are scaled up by the expansion factor.

This gel transfer should be carried out in a timely manner to avoid dehydration of the gel: we recommend finishing this step within 5 min.

11a. After the mounting, immediately add a small amount of water (or diluted PBS for ExFISH samples) to keep the gel hydrated and avoid gel shrinkage.

Sample mounting with agarose

5b. Coverslips, glass-bottom dishes, and glass-bottom multi-well plates can be used as sample holders. Slides or plastic multi-well plates can also be used, but the thickness of the sample holder will limit the choice of objectives and/or cause aberrations on inverted microscopes.

6b. Prepare 0.5% (w/v) low-melting agarose solution in the correct imaging medium. For example, for proExM samples, prepare 0.5% (w/v) agarose in water. For ExFISH samples, prepare 0.5% (w/v) agarose in diluted PBS or other buffers of your choice. Keep the agarose solution warm in a 40°C water bath to avoid premature hardening. If hardened, use a short microwave cycle (10 to 20 sec) to melt the agarose solution. Alternatively, heating it to ∼60°C will re-melt it as well.

7b. Transfer the expanded gel to the sample holder of choice (e.g., coverslips and glass-bottom dishes). During the transfer, remove and wick away liquid from the expanded gel (see step 10a, above).

8b. Using a pipet, carefully apply the melted agarose solution onto the edges of the gel. If necessary, you can also embed the entire gel in agarose, using the melted agarose solution.

9b. Wait for the agarose to harden at room temperature or at 4°C, and then add a small amount of water (or diluted PBS for ExFISH samples) to prevent dehydration.

10b. After imaging is complete, you can place the mounted sample in high-salt buffer such as PBS. This will shrink the gel but not the agarose. Using paintbrushes or forceps, you can separate the gel from the agarose. The gel can be then stored in PBS for later use.

Sample mounting with superglue

5c. Coverslips, glass-bottom dishes and glass-bottom multi-well plates can be used as sample holders. Slides or plastic multi-well plates can also be used. However, since the superglue turns opaque upon curing, this mounting method is not compatible with inverted microscopes that require imaging through the sample holder.

6c. Clean the surface of the sample holder by rinsing with water and then ethanol. After rinsing, let the holder dry in air.

7c. Apply superglue to the surface of the sample holder. Only apply the glue to the area where the gel should be attached. Also make sure excessive superglue is removed by paper wipes, leaving only a thin layer.

8c. Transfer the expanded gel to the sample holder of choice. Before the transfer, remove and wick away liquid from the expanded gel (see step 10a).

9c. Once the gel is placed on the glue, wait for 20 to 30 sec for the superglue to cure before adding water (or diluted PBS for ExFISH samples) to hydrate the gel.

After curing, there should be an opaque interface between the gel and the superglue.

Imaging ExM samples with inverted microscopes (e.g., inverted spinning disk confocal microscopes)

A typical inverted microscope setup is shown in Figure 10. The poly- L -lysine mounting method is recommended for inverted microscopes, since the transparent interface between the gel and the poly- L -lysine modified surface will allow for imaging through the sample holder.

For imaging on an inverted microscope, we recommend starting with low-magnification (4× or 10×) objectives to find the region of interest first, and then switching to high-magnification objectives. Air objectives are typically low-magnification objectives (e.g., 4× or 10×) and can be used at the beginning to find the samples and the regions of interest. For high-magnification imaging (>20×), we recommend using water-immersion objectives to avoid aberrations caused by refractive index mismatch between the aqueous mounting medium and the coverslip or oil. Due to the generally short working distances, oil-immersion objectives can be considered for use mainly when the expanded gel is mounted close (within ∼10 μm) to the coverslip and when the specimen or object of interest is close to the objective (such as in cultured cells or thin tissues). The working distance of the objective is typically not an issue for thin samples such as cultured cells, if the gel is properly mounted on the sample holder (i.e., the sample side of the gel is mounted facing the objective side using the poly- L -lysine method). For some tissue samples, however, the working distance becomes an issue when imaging deep into the gel (e.g., >300 µm). In this case, long-working distance water-immersion objectives are recommended. As with every microscopy experiment, make sure the correction collar on the objective (if applicable) is adjusted to match the thickness of the coverslips.

Many of the wide-field and confocal microscopes used in biology labs and imaging facilities are inverted microscopes. We have found that spinning-disk confocal microscopes offer a good combination of resolution and speed, and are highly suitable for fast nanoscale imaging of expanded tissues. Laser-scanning confocal microscopes can be used most effectively for high signal-to-background imaging of thin and small samples such as cultured cells.

Imaging with upright microscopes

For upright microscopes, any of the three mounting methods, poly- L -lysine, agarose, or superglue, can be used. If you plan to retrieve your sample after imaging, we recommend the agarose mounting method. In the other two methods, the gel is physically attached to the sample holder, but can potentially be cut free using a razor blade if the sample needs to be retrieved (note that this process can damage the sample, if you are not careful). For long-term imaging (e.g., over a few days), we recommend using the superglue mounting method, as it offers the strongest adhesion to the sample holder. The poly- L -lysine method gives a transparent interface between the gel and the sample holder and thus can be imaged on an inverted microscope, too. For imaging on an upright microscope, we recommend starting with low-magnification (4× or 10×) objectives to find the region of interest first, and then switching to high-magnification objectives, which is similar to the procedure on an inverted microscope. For high-magnification imaging (>20×), we recommend using water-dipping or water-immersion objectives, as the gel is immersed in aqueous mounting medium.

Imaging with microscopes with the sample hanging from above (e.g., light-sheet microscopes)

Light-sheet microscopes are suitable for imaging of expanded tissue samples at high speeds, using long-working-distance objectives. To accommodate the objectives, some light-sheet microscopes hang the samples from above. For example, a typical imaging setup of a light-sheet microscope (Zeiss Z.1 light-sheet microscope) is shown in Figure 11. The Z.1 light-sheet microscope uses a rod to hang samples from above the objectives, perpendicular to the microscope's light path. The sample is then enclosed in an imaging chamber filled with aqueous mounting medium. For expanded ExM samples, a thin coverslip can be used as the backing to physically support the gel, and then the coverslip backing can be attached to the rod through a small 3-D-printed adapter, which is available for download at our Web resource http://expansionmicroscopy.org. To attach the gel to the coverslip backing, we recommend the superglue mounting method, as it most strongly attaches the gel to the coverslip. Poly- L -lysine mounting can be also used for short-term imaging, but we have observed detachment of the gel if the mounting is not properly performed (e.g., due to expired poly- L -lysine, fast stage movements, or inadequate water removal from the gel before placement). After mounting the gel to the coverslip backing, the coverslip backing can be attached to the adapter with superglue. The adapter is designed to match the dents at the end of the rod so it can be mechanically fixed to the rod.

On the Zeiss Z.1 light-sheet microscope, we typically use a 1.0 NA, 20× long-working-distance water-immersion objective for high-resolution imaging. If the imaging volume is large and requires a long imaging time, we can use lower-magnification objectives (5×) to obtain a low-resolution, quick overview of the sample. The Z.1 light-sheet microscope is suitable for imaging large, multi-color samples with its two-camera setup, fast image acquisition, and capability of capturing two color channels simultaneously. In addition, the long working distance of the objective can be used to image large transparent samples. These specifications of the Z.1 light-sheet microscope are particularly advantageous for imaging ExM samples.

We first demonstrated these advantages by imaging a ∼575 × 575 × 160–µm volume (∼6 × 10 10 voxels in three colors) of a HCR-ExFISH sample in about ∼3 hr, with a resolution capable of resolving single RNA puncta within the expanded tissue (Fig. 11D-F Chen et al., 2016 ). Even larger samples in the millimeter range have been successfully imaged with this system. One such example is shown in Figure 11G, in which a subregion of a mouse hippocampus infected with a Brainbow 3.0 AAV virus (Fig. 5) was expanded with a proExM protocol described earlier. The imaging volume was ∼1.5 × 0.8 × 0.1 mm in pre-expansion units, and the dataset required stitching of 180 individual 3-D image stacks, resulting in a 5 Terabyte, 0.7 Teravoxel dataset with an effective lateral pixel size of about 60 nm.


Lysate Preparation: Why is RIPA Buffer Best for Western Blot?

In 1979, Jaime Renart et al. published an article entitled “ Transfer of proteins from gels to diazobenzyloxymethyl -paper and detection with antisera: a method for studying antibody specificity and antigen structure,” the prelude to the modern Western blot (WB) technique.

Soon after, Harry Towbin et al. went one step further and published “Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications.” With that, the WB technique was officially born.

Today, WB experiments are a cornerstone of biological research. Unfortunately, it is frequently a challenge to obtain good results.

A wise man once said:

A successful WB relies upon:

Speaking of W B, as an original manufacturer of all our products, Proteintech’s R&D staff test on average over 70 samples for each product in WB. Proteintech’s senior R&D staff will share with you our years of experience with WB to ensure every WB is a successful one.

What Goes into a Lysis Solution?

A lysis solution contains the following components:

1. The buffer system

The pH of the solution is critical. Proteins may precipitate or become unstable when the pH is outside of the physiological range. To avoid this situation, a buffer system such as Tris-HCl is recommended. Besides buffering solutions in this range, a Tris-HCl buffer preserves the physiological ionic strength and prevents the formation of insoluble products with other ions. A HEPES buffering system is another option. We recommend avoiding buffers with high concentrations of potassium, because these can precipitate proteins when sodium dodecyl sulfate (SDS) is present.

2. Salt ions

When the salt ion concentration is too high, some proteins may precipitate. Additionally, when ion concentration is too high, a “smiley face” of band migration may result.

3. Chaotropic agents

Chaotropic agents weaken the hydrophobicity of the proteins to solubilize them. There are two kinds of chaotropic agents in a lysis buffer:

a. Urea/thiourea. These molecules unravel hydrophobic regions by disrupting hydrogen bonding between amino acids. Usually when doing protein extraction for a WB, 6­–8M urea and/ or 2M thiourea can be used.

b. Detergents. These are a broad class of surfactants. The key to their solubilizing power is their amphiphilic structure. The hydrophobic end binds to the hydrophobic portions of proteins while the hydrophilic end interacts with water, resulting in solubilization.

Ionic detergents can be further divided into cationic, anionic, and amphoteric detergents. Common ionic surfactants are SDS, DOC (sodium deoxycholate), and SLS (sodium lauryl sarcosine). Common amphoteric surfactants are CHAPS ( 3-[(3-Cholamidopropyl)dimethylammonio]-1-propanesulfonate) , and CHAPSO ( 3-[(3-cholamidopropyl)dimethylammonio]-2-hydroxy-1-propanesulfonate) . Common non-ionic surfactants are Triton X-100, Triton X-114, Tween-20, and NP40.

Note: It is critical in a WB that the number of negatively charged SDS molecules that bind to a protein is proportional to the protein’s mass, so that the migration rate is influenced only by mass. Adding cationic surfactants in the lysis buffer would disrupt the SDS-protein interaction and make the proteins migrate in the opposite direction.

Due to the complexity of protein biochemistry, it is challenging to predict the optimal surfactant to extract a given protein. Thus, experimenting with different surfactants is recommended if you encounter issues. This is especially recommended for membrane proteins.

Related: Prestained Protein Ladders
Protein Ladder (Free sample available) Broad Range Protein Ladder
Catalog no . : PL00001 Catalog no. : PL00002
Molecular weight coverage : 10-180 kDa Molecular weight coverage : 3-245 kDa
No. of markers : 10 prestained proteins No. of markers : 13 prestained proteins
Resolution of the prestained protein ladder in a 10-20% Tris-glycine gel (SDS-PAGE). Resolution of the prestained protein ladder in a 4-12% Bis-Tris gel (SDS-PAGE)
4. Protease inhibitors

Tissues and cells often contain large amounts of proteases. During lysis, these are released and, in turn, can digest the target protein. Therefore, protease inhibitors are critical for preserving the target protein. Common protease inhibitors are PMSF (phenylmethylsulfonyl fluoride), Aprotinin, Leupeptin, Pepstatin, and AEBSF-HCL (4-benzenesulfonyl fluoride hydrochloride). PMSF is highly effective and is the most popular choice for lysate preparation.

Many protease inhibitors require a divalent metal ion to function, so a sequestering agent is also often used to inhibit protease activity, such as EDTA. In addition, for phosphorylated target proteins, a phosphatase inhibitor such as sodium fluoride or sodium orthovanadate is needed to preserve the phosphorylated form of the protein. Sodium orthovanadate, in particular, is very effective, but needs to be activated by adjusting the pH of the solution to 10 and then boiling it until the solution is colorless. Other phosphatase inhibitors include sodium pyrophosphate and β-glycerol phosphate.

5. Reductant

Many proteins exist in multimers through disulfide bonds. Reductants disrupt these bonds so that the extracted proteins are present in monomeric form. Common reducing agents are DTT (dithiothreitol) and BME (beta-mercaptoethanol). Keeping all of this in mind, RIPA buffer is the best choice for sample lysate preparation. We have validated over 13,000 antibodies in WB, and time and time again, experience the best results using RIPA buffer. Over the years we have refined the buffer and below you will find Proteintech’s optimized version:

RIPA buffer For 1000ml
50mM Tris HCl, PH 7.4 50ml
150mM NaCl 8.76ml
1% Triton X-100 or NP-40 10ml
0.5% Sodium deoxylcholate 5g
0.1% SDS 1g
1mM EDTA (0.5M stock) 2ml
10mM Naf 0.42g
Add ddH2 O to 1000ml
Add PMSF to a final concentration of 1mM and any other protease inhibitors immediately before use.
4 x SDS sample buffer For 1000ml
12% SDS 120g
25% Glycerol 250ml
150 mM Tris.HCl (pH 7.0 1M stock) 150ml
0.03% Bromophenol Blue 300mg
20% β-mercaptoethanol 200ml
Add ddH2O to 50ml, aliquot and store at -20 °C
20% β-mercaptoethanol , (or 500 mM DTT replaced), should be added freshly before use.

If you are experiencing issues extracting the conventional lysate protein, we recommend reading the literature, rather than randomly trying various extraction kits.

Related: Loading control antibodies

GAPDH is commonly used as a protein loading control in western blot due to its consistently high expression in most cell types. This enzyme participates in several cellular events such as glycolysis, DNA repair, and apoptosis.

Proteintech monoclonal GAPDH antibodies are raised against a whole-protein antigen of human origin and have over 2,670 citations .

Beta-actin is usually used as a loading control due to its broad and consistent expression across all eukaryotic cell types and the fact that expression levels of this protein are not affected by most experimental treatments.

66009-1-Ig has been cited over 1,135 publications and has wide species reactivity.

Proteintech control antibodies are just $99 each ( € 150 in Europe) for a 150ul size vial.


Conclusions

The targeting of newly synthesized MCF proteins is obviously a matter of consecutive steps: A first decision is already attained in binding to distinct chaperone proteins. If a newly synthesized protein binds to Pex19p, it can be targeted to peroxisomes as demonstrated by Ant1p. Hsp70-bound preproteins lacking a high affinity Pex19p interaction site can get into contact with mitochondrial Tom70 and bind at the site marked by cysteine C141. However, their subsequent fate is mainly determined by interactions with Tom40 that serves both as a general import pore and as a general selectivity filter: MCF proteins exposing a positively charged matrix loop, as well as preproteins containing a positively charged presequence, will be recognized by Tom40 and insert into the import pore. Tom70-bound proteins containing a negatively charged loop, such as Ugo1, may be expelled from the Tom40 import channel and insert into the lipid phase of the mitochondrial outer membrane, other proteins may be released and stay in the cytosol. Further studies on the selection and translocation of carrier proteins should be facilitated by the new data on the high-resolution structures of the import machinery that have recently become available, and by the identification of the bipartite MCF targeting sequence.


What is the epithelial to mesenchymal transition (EMT)?

Epithelial to mesenchymal transition (EMT) is the process whereby epithelial cells are transformed into mesenchymal cells. Epithelial cells form the epithelium tissue which covers the internal and external body surface of an organism. These cells are polarized and form extensive cell-cell adhesions, including adherens junctions and tight junctions, with each other [1] [2] . One of the fundamental functions of epithelial tissue is to act as a protective barrier for the underlying tissue and organs. Mesenchymal cells are, however, not well organized in a three-dimensional extracellular matrix [3] . During development, mesenchymal cells can gain the ability to migrate and differentiate into other cell types. This gives rise to the formation of defined structures, such as the neural crest [3] .

During EMT, epithelial cells lose their polarity, as well as their cell-cell adhesions, and gain the ability to migrate, proliferate, differentiate and develop into specific tissues and organs. EMT is classified into 3 main types:

  • type 1 EMT is associated with development such as embryo formation, and organ development
  • type 2 EMT is associated with repair processes such as wound healing, tissue regeneration, and organ fibrosis
  • type 3 EMT is associated with tumor metastasis [3][4] .

Here we will only consider the EMT in development where it plays important roles in producing the various tissues and organs, and defining their arrangement in the body.

Stiff matrix promotes integrin clustering which promotes the recruitment of Rac1b to the membrane, facilitating its interaction with NADPH oxidase and the subsequent production of ROCs which promote Snail expression and hence EMT. On the other hand, Rac1b is less likely to be recruited to the membrane of cells in a softer matrix. EMT is less likely to occur in this situation.

Importantly, EMT is reversible, meaning a mesenchymal to epithelial transition, (MET) can occur. In fact, several cycles of EMT and MET are required before the final architecture of the organ is established. These rounds of EMT are known as primary, secondary and tertiary EMT [3] . An example of primary EMT occurs during gastrulation, where embryonic epithelium undergoes EMT to form the mesoderm. In vertebrates, gastrulation is induced by proteins from the transforming growth factor &beta (TGF&beta) superfamily, especially Nodal and Vg1 [5] [6] . These TGF&beta proteins, together with the aid of fibroblast growth factor (FGFs) signaling, induce the Snail gene [7] [8] [9] , which is a key transcription factor for EMT [10] . Snail promotes the breaking away of the epithelial cells from the basal membrane prior to their EMT, by repressing E-cadherin expression [11] and activating metalloproteinases [8] . This cleaves the cell adhesion complexes and degrades extracellular matrix, respectively, to aid cell migration [10] . Nodal and Vg1 also promotes Wnt signalling [6] [12] , which co-operates with FGF and Snail to orchestrate EMT [9] .

Physical properties regulate EMT

The molecular mechanisms that drive EMT are complex and involve many more biochemical components. These have been reviewed extensively in [10] . In addition to the biochemical signals, mechanical signals, such as the rigidity of the cell&rsquos microenvironment, also play roles in the regulation of EMT. Lee et al. observed that mammary epithelial cells underwent EMT when placed on hard substrates and treated with matrix metalloproteinase-3 (MMP3). This was not observed in cells grown on soft substrates [13] . Furthermore, treatment with MMP3, caused cells on both substrates to increase their expression of Rac1b. However, only those cells on the hard substrate produced reactive oxygen species (ROS) [13] which is downstream of MMP3 and Rac1b [14] . Interestingly, in cells on hard substrates, Rac1b is recruited to the cell membrane through integrin clustering. This allows Rac1b to assemble with NADPH oxidase and contribute to ROS production. It was determined therefore, that cells growing on rigid substrates will undergo EMT after the expression of Snail is increased following Rac1b mediated ROS stimulation [14] . Stiffer substrates have been shown to promote integrin clustering [14] and focal adhesion formation [15] , and it is this process that leads to Rac1b recruitment and its interaction with NADPH oxidase that produces ROS, which in turn promotes EMT. As soft substrates do not promote integrin clustering, this cascade of events does not occur, and cells growing on soft substrates do not undergo EMT.

Besides the rigidity of the cell microenvironment, other mechanical stresses placed on epithelial cells from their neighbouring cells, are implicated in the initiation of EMT. This was described by Gomez et al. who used microfabrication techniques to generate a confluent, square-shaped sheet of mammary epithelial cells. After treating them with TGF&beta (an inducer of EMT), the team observed that the cells underwent EMT preferentially at the corner and edge of the square, which correspond to regions of higher mechanical stress [16] . High levels of myocardin-related transcription factors A (MRTFA) was also found to be localized in the nucleus of cells located at regions of higher mechanical strain [16] .

When mechanical strain is lower, MRTFA binds to actin monomers and is retained in the cytoplasm [17] . However, when mechanical stress increases, the tension of cytoskeleton increases and this activates the Rho-actin signaling pathway, which reduces the amount of monomeric actin in the cytoplasm, causing MRTFA to translocate to the nucleus [17] [18] . Here, MRTFA forms a complex with Smad3, which binds to the promoter region of the SNAI2 gene, activating its transcription and subsequently dissolving cell-cell contacts [19] . MRTFA is a co-activator of serum response factor (SRF) which upregulates actin cytoskeletal proteins leading to cytoskeleton remodelling [19] . This dual pathway is activated following the nuclear localization of MRTFA, and promotes epithelial cells to undergo EMT.

The direct application of biomechanical force has also been found to be sufficient to induce EMT. Zhou et al. stretched human keratinocytes by applying a 10% elongation to these cells, and observed that the applied force can directly activate NF-&kappaB, which in turn promotes EMT [20] [21] [22] . Similarly, the application of compressive force to HEK-293 cells was shown to induce the expression of EMT regulators SNAI2 and ZEB1, along with the messenchymal marker VIM, via the RhoA/ROCK pathway [23] . Taken together, these experimental results highlight that underlying EMT is a complex network of signalling that involves not only biochemical signals, but also a variety of mechanical cues.


How the Bradford Protein Assay Works

The Bradford protein assay is a time-tested colorimetric assay. When the Bradford reagent (acidified Coomassie Brilliant Blue G-250) binds to proteins, the dye undergoes a color change in the visible spectrum, with the absorbance maximum moving from 470 to 595 nm. The absorbance at 595 nm is then read either in a spectrophotometer or a microplate reader and is directly proportional to the amount of protein bound. The exact protein concentration of the sample is determined by interpolation from a standard curve made by measuring the absorbance of a dilution series of protein standards of known concentrations within the linear response range of the Bradford protein assay. Proteins commonly used as standards include bovine serum albumin (BSA) and bovine &gamma-globulin (BGG).

There are two types of Bradford protein assay kits. One contains ready-to-use reagent and prediluted BSA or BGG standards for a simple and quick Bradford protein assay. The other kit provides more flexibility with 5x concentrated reagent and lyophilized BSA or bovine-globulin standards, allowing the user to prepare reagent and standards at any concentration.


UV cross-linking and immunoprecipitation (CLIP)

Interest in RNA-protein interactions is booming as we begin to appreciate the role of RNA, not just in well-established processes such as transcription, splicing, and translation, but also in newer fields such as RNA interference and gene regulation by non-coding RNAs.

CLIP is an antibody-based technique used to study RNA-protein interactions related to RNA immunoprecipitation (RIP) but differs from RIP in the use of UV radiation to cross-link RNA binding proteins to the RNA that they are bound to. This covalent bond is irreversible, allowing stringent purification conditions. Unlike RIP, CLIP provides information about the actual protein binding site on the RNA.

Different types of CLIP exist, high-throughput sequencing-CLIP (HITS-CLIP), photoactivatable-ribonucleoside enhanced CLIP (PAR-CLIP), and individual CLIP (iCLIP).

Here is a summary of the iCLIP protocol adapted from Konig et al. J. Vis. Exp. 2011. “iCLIP -Transcriptome-wide Mapping of Protein-RNA Interactions with Individual Nucleotide Resolution.”

Further adapted from Huppertz et al. Methods. 2014. “iCLIP: Protein–RNA interactions at nucleotide resolution.”

Lysis buffer

Protease inhibitors (add fresh each time)

High-salt Wash

Low RNase dilution High RNase dilution

1/500 RNase I dilutions for library preparation 1/50 RNase I dilutions to control for antibody

Wash buffer PNK mix

4 μL 5x PNK pH 6.5 buffer [350 mM Tris-HCl, pH 6.5 50 mM MgCl 2 5 mM dithiothreitol]

Ligation mix A Hot PNK mix

4 μL 4x ligation buffer [200 mM Tris-HCl 40 mM MgCl 2 4 mM dithiothreitol]

1.5 μL pre-adenylated linker L3 [20 μM]

PK buffer PKurea buffer

RNA/primer mix RT mix

0.5 μL Rclip primer [0.5 pmol/μL]

0.25 μL Superscript III reverse transcriptase

Ligation mix B Oligo annealing mix

0.8 μL 10x CircLigase Buffer II

1 μL primer mix P5/P3 solexa

20 μl Accuprime Supermix 1 enzyme

​ 1. UV cross linking of tissue culture cells

1.1. Remove media and add ice-cold PBS to cells (eg use cells grown in a 10 cm plate for three experiments and add 6 ml PBS).

1.2. Remove lid, place on ice and irradiate once with 150 mJ/cm 2 at 254 nm using a Stratalinker.

One or more negative controls should be maintained throughout the complete experiment. Knockout cells or tissue, as well as non-cross-linked cells, are good negative controls, while knockdown cells are not recommended.

Note. Optional 4-thiouridine pre-incubation and UV-A crosslinking could be used for certain proteins. 4-thiouridine enhances crosslinking of some proteins. (For this optional step follow the instructions below).

Optional – Thiouridine labeling and UV-A crosslinking (alternative to step 1.2)

• Add 50 μL of 4SU (stock concentration: 100 mM 4SU) to a 10 cm plate of cells cultured in 10 ml DMEM supplemented with FBS and pen strep to get a final concentration of 500 μM 4SU. Alternatively, add 10 μL of 4-thiouridine (stock concentration: 100 mM) to get a final concentration of 100 μM 4SU.

• Incubate cells with 4-thiouridine for 60 min at 500 μM or 8 h at 100 mM. Check cell viability.

• Aspirate the medium and add 6 mL of ice-cold PBS to cells growing in a 10 cm plate (enough for three immunoprecipitations). Remove the lid and place on ice.

• Irradiate once with 2 × 400 mJ/cm 2 in a Stratalinker 2400 with 365 nm bulbs, or an equivalent instrument.

1.3. Harvest cells with a cell scraper and transfer cell suspension to microtubes (eg 2 mL to each of three microtubes).

1.4. pellet cells (spin at top speed for 10 sec at 4°C), then remove supernatant.

1.5. Snap-freeze cell pellets on dry ice and store at -80°C until use.

The experiment could take up to a week. Avoid multiple cycles of freeze thaw.

1.6 Bead preparation

1.6.1 Add protein A or protein G beads (e.g. 100 μl magnetic beads per experiment) to a fresh microtube and wash beads 2x with lysis buffer.

1.6.2 Resuspend beads in lysis buffer (100 μL), add antibody (2-10 μg) and rotate tubes at room temperature for 30–60 min.

The amount of antibody required might need to be optimized. A no-antibody sample is a good negative control.

1.6.3. Wash beads 3x with lysis buffer (900 μL) and leave in the last wash until ready to proceed with the immunoprecipitation (step 3.1).

If an antibody is working in IP, this is a good indication that it will work in CLIP.

2. Cell lysis and partail RNA digestion

2.1. Resuspend cell pellet in lysis buffer with protease inhibitors (1 mL) and transfer to 1.5 mL microtubes.

Note. Here you can try an optional sonication step. Sonication shears the DNA, which releases proteins from the chromatin increasing recovery of protein–RNA complexes.

Optional – Sonication of samples

Sonicate the sample on ice using a probe sonicator. The probe should not touch the tube sides to avoid foaming. Sonicate twice with 10 sec bursts at five decibels. Clean the probe by sonicating H 2 O before and after sample treatment.

Use Bioruptor plus for five cycles with alternating 30 sec on/30 sec off at low intensity.

2.2. Add low RNase dilution (10 μL) and Turbo DNase (2 μL) to the cell lysate and incubate for exactly 3 min at 37°C, shaking at 1,100 rpm, then immediately transfer to ice.

2.3. Spin at 4°C at 22,000 g for 10 min and carefully collect the cleared supernatant (leave about 50 μL lysate with the pellet).

Each member of the laboratory should use their own set of buffers and reagents to easier identify potential sources of contamination. Ideal conditions for the RNase digestions may need to be optimized for every new batch of RNase.

3. Immunoprecipitation and dephosphorylation of RNA 3' ends

3.1. Remove lysis buffer from the beads (step 1.1.3) and add cell lysate (from step 2.3).

3.2. Rotate the samples for 2 h at 4°C.

3.3. Discard the supernatant, wash beads 2x with high-salt buffer (900 μL) and then 2x with wash buffer (900 μL). Rotate these washes for at least 1 minute at 4°C.

Optimization and stringent washing conditions are very important.

Note. In the case that standard iCLIP conditions are not stringent enough to specifically purify the protein of interest, a dual immunoprecipitation with urea denaturation can be used instead of the protocol described here. For more information see Huppertz et al. Methods. 2014. “iCLIP: Protein–RNA interactions at nucleotide resolution.”

3.4. Discard the supernatant, resuspend beads in PNK mix (20 μL) and incubate for 20 min at 37°C in a thermomixer.

3.5. Add wash buffer (500 μL), wash 1x with high-salt buffer and then 2x with wash buffer.

4. Linker ligation to RNA 3' ends and RNA 5' end labeling

4.1. Carefully remove the supernatant, resuspend beads in ligation mix A (20 μL) and incubate overnight at 16°C in a thermomixer.

4.2. Add wash buffer (500 μL), wash 2x with high-salt buffer (1 mL) and then 2x with wash buffer (1 mL). Carry out these washes with rotation at 4°C for 5 mins each.

4.3. Remove the supernatant, resuspend beads in hot PNK mix (4 μL) and incubate for 5 min at 37°C.

4.4. Remove the hot PNK mix and resuspend beads in 1x SDS-PAGE loading buffer (20 μL).

4.5. Incubate on a thermomixer at 70°C for 10 min.

4.6. Immediately place on a magnet to precipitate the empty beads and load the supernatant on the gel (see step 5).

5. Linker SDS-PAGE and membrane transfer

5.1. Load samples as well as a pre-stained protein size marker (5 μL) on a precast 4-12% Bis-Tris gel, and run the gel for 50 min at 180 V in 1x MOPS running buffer (according to manufacturer's instructions).

Gels with constant pH 7 are recommended.

5.2. Remove the gel front and discard as solid waste (contains free radioactive ATP).

5.3. Transfer the protein-RNA complexes from the gel to a nitrocellulose membrane using a wet transfer apparatus (transfer 1 h at 30 V depending on manufacturer's instructions).

5.4. Following the transfer, rinse the membrane in PBS buffer, then wrap it in clingfilm and expose it to a film at -80°C for 30 min, 1h and then overnight.

A fluorescent sticker next to the membrane later allows to align the film and the membrane. The success of the experiment can be monitored at the autoradiograph of the protein-RNA complex after membrane transfer.


Final words

In this chapter (Enzymes Involved in DNA Replication) and the next, we will examine the process of replication.

After describing the basic mechanism of DNA replication, we discuss the various techniques researchers have used to achieve a complete understanding of replication.

Indeed, a theme of this chapter is the combination of genetic and biochemical approaches that have allowed us to uncover the mechanism and physiology of DNA replication.