3D Imaging 1: Principles of FISSEQ + Expansion Microscopy

15th of November - Evan R. Daugharthy (ReadCoor, Harvard) & Paul Reginato (MIT)

Recordings

• Review 09: Synthetic Development Biology
• Lesson 10: Principles of FISSEQ + Expansion Microscopy

Presentations and slides

• 9:00 Class starts

Introduction

Why do we need analytic tools for synthetic projects? The tools for synthetic biology have grown incredibly powerful: DNA synthesis, genome engineering, synthetic cells, directed evolution, cell-free systems, metabolic engineering, and nanomaterial science. However, these tools only cover the second half of the "read/write" cycle. In this class, we will discuss the rationale for developing measurement technologies ("read") to complement these engineering tools ("write"), so that we can understand the effects of our bioengineering efforts and make new products that resemble real biological systems.

We will review various approaches to molecular measurements, including DNA and RNA sequencing, proteomics, imaging 3D structural morphometry. We will focus predominantly on in situ detection of single molecules (in situ is latin for "in place," referring to detection of molecules in their native spatial arrangement in cells/tissues). Finally, we will discuss applications of these technologies to fibroblast wound healing, understanding how the brain works, and to developing new organoids to further our understanding of biological development and create new biomedical interventions to advance human health.

Readings

Background Reading:

The FISSEQ Method: Lee J, Daugharthy E, Scheiman J, Kalhor R, Yang JL, Ferrente TC, Terry R, Jeanty SSF, Li C,Amamoto R, Peters DT, Turczyk BM, Marblestone A, Inverso S, Bernard A, Mali P, Rios X, Aach J, Church GM (2014) [Highly multiplexed three-dimensional subcellular transcriptome sequencing in situ] (http://arep.med.harvard.edu/pdf/Lee_Sci_2014.pdf). Science 343(6177):1360-3.

Expansion Microscopy: Chen et al. (2015) [Expansion microscopy] (http://syntheticneurobiology.org/PDFs/15.01.chen.FULL.pdf). Science 347(6221): 543-548

Theory of RNA and Cellular Molecular State: Kim, Junhyong, and James Eberwine (2010) [RNA: state memory and mediator of cellular phenotype] (http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2892202). Trends in cell biology 20.6:311-318.

Additional Reviews of FISSEQ and Single-Cell Sequencing

Ginart, Paul, and Arjun Raj (2014) [RNA sequencing in situ] (http://rdcu.be/eoW7). Nature biotechnology 32.6:543-544.

Mignardi, Marco, and Mats Nilsson (2014) [Fourth-generation sequencing in the cell and the clinic] (http://www.genomemedicine.com/content/6/4/31). Genome medicine 6.4:31.

Avital, Gal, Tamar Hashimshony, and Itai Yanai (2014) [Seeing is believing: new methods for in situ single-cell transcriptomics] (http://www.genomebiology.com/2014/15/3/110). Genome biology 15.3:110.

Additional methods for Expansion Microscopy

Chen et al. (2016) [Nanoscale Imaging of RNA with Expansion Microscopy.] (http://syntheticneurobiology.org/PDFs/16.07.chen.FULL.pdf). Nature Methods 13.8:679–684

Tillberg et al. (2016) [Protein-Retention Expansion Microscopy of Cells and Tissues Labeled Using Standard Fluorescent Proteins and Antibodies] (http://syntheticneurobiology.org/PDFs/16.07.tillberg.FULL.pdf) Nature Biotechnology 34.9: 987–992

Chang et al. (2017) [Iterative expansion microscopy.] (http://syntheticneurobiology.org/PDFs/17.04.chang.FULL.pdf) Nature Methods 14.6:593–599

Additional Theory

Eberwine, James, and Junhyong Kim (2015) [Cellular Deconstruction: Finding Meaning in Individual Cell Variation] (http://www.sciencedirect.com/science/article/pii/S0962892415001282). Trends in cell biology 25.10:569-578.

Trapnell, Cole (2015) [Defining cell types and states with single-cell genomics] (http://genome.cshlp.org/content/25/10/1491.full). Genome research 25.10:1491-1498.

Additional Historical Context of DNA Sequencing

Hutchison, Clyde A. (2007) [DNA sequencing: bench to bedside and beyond.] (./files/nucl.acids_res.-2007-hutchison-6227-37.pdf) _Nucleic acids research 35.18: 6227-6237.

Goodwin, Sara, John D. McPherson, and W. Richard McCombie. (2016) [Coming of age: ten years of next-generation sequencing technologies.] (./files/nrg.2016.49.pdf) Nature Reviews Genetics 17.6: 333-351.

Additional Background on Super-resolution Imaging

Sydor et al. (2015) [Super-Resolution Microscopy: From Single Molecules to Supramolecular Assemblies] (http://www.cell.com/trends/cell-biology/pdf/S0962-8924(15)00191-9.pdf) Trends in Cell Biology 25.12:730-748

Class Assignments

Reagents and tools needed for ExM homework:

Exercise 1: Cast an expanding gel

WARNING: Acrylamide and other reagents in this procedure are hazardous in case of skin contact (permeator), of eye contact (irritant), of ingestion, of inhalation, and are also known carcinogens. Always read and follow the MSDS (Material Safety Data Sheet), use PPE (Personal Protective Equipment), and follow all necessary safety procedures!

Reagents:

Note 1: Sodium Acrylate sometimes comes with a variable purity level, which can affect performance. For every fresh bottle purchased, we make a 38g/100mL (33 wt%) sodium acrylate stock and check that it is colorless under normal room lighting. If the stock has a yellow tint, we discard the bottle from which it was made. For the purposes of this assignment, however, it will work well enough that you should proceed even if your sodium acrylate solution has a yellow tint.

Note 2: Once open, we have been storing sodium acrylate in an airtight, low humidity, or dessicator chamber in -20 degrees C, since the solid is moisture sensitive. APS powder and 100% TEMED solution are stored in a room temperature dessicator. We have been storing the monomer solution mixed up at -20 degrees C for up to 1 month. TEMED and APS solutions can be kept in -20 degrees C, and we generally remake the TEMED and APS stocks at least once every 2 weeks.

Protocol:

Step 1. Prepare the Monomer Solution (can be stored at -20 for at least a month):

Note 1: If you are preparing your own sodium chloride stock solution rather than buying it in solution, you will have trouble getting it to dissolve up to 5M. In that case, use 4M sodium chloride and 49.75 uL of it, and add 0 uL of water so that the final volume is the same.

Note 2: Right before gelation, 2 ul each of 10% TEMED and 10% APS will be added, bringing the final volume up to 100 ul (see below).

Step 2. Prepare gelation chamber: Obtain two glass slides (~ 3"x1"), two coverslips (~ 22mmx22mm), and some parafilm. Wrap one of the slides in parafilm such that one face of the slide has a smooth flat surface one parafilm layer thick. (The other face of the slide will have the folded edges of the parafilm on it.) Press the parafilm on the flat side to ensure there is no gap between the parafilm and the slide. Place a coverslip at each end of the parafilm-wrapped slide on its smooth face. Place this parafilm-wrapped slide with coverslips into a petri dish or any container with a lid. Set the remaining glass slide aside.

Step 3. Prepare 10% (v/v) TEMED and 10% (w/v) APS. 10% TEMED can be made by simply diluting pure TEMED tenfold in water. 10% APS can be made by measuring out some APS, finding its weight in milligrams, multiplying the weight by 9.5, and then adding that many microliters of water. Eg for 30 mg of APS, add 285 ul of water. (This assumes that 10% (w/w) APS is ~ 5% (v/w), which is approximately correct.)

Step 4. Prepare Gelation Solution: Mix the following 4 solutions on ice: Monomer solution, 10% TEMED (accelerator), 10% APS (initiator solution). (APS initiator solution needs to be added last to prevent premature gelation). Solutions should immediately be vortexed or pipetted up and down after mixing to ensure full mixing and the gel should then immediately be cast to prevent premature gelation. For 50 µL gelling solution, mix the following: a) Monomer solution (48µl) b) Accelerator solution (1µl): 10% TEMED (TEMED stock solution at 10%, final concentration 0.2% (w/w). (Accelerates radical generation by APS). c) Initiator solution (1µl): APS (APS stock at 10%, final concentration 0.2% (w/w)). (This initiates the gelling process. This needs to be added last).

Step 5. Polymerize the Gel: Pipet 10 ul of gelation solution onto the flat surface of the parafilm-wrapped slide between the coverslips and place the other glass slide on top so that it rests on the coverslips, leaving a small space between the two slides where the gel will polymerize. Put a wet kim-wipe or tissue in the container to keep the air in the container hydrated. Put the lid on the container and place it in a 37C incubator for 1 hour (or 1.5 hours at room temperature) to polymerize. (Note that the most ideal condition for polymerization is oxygen-free because oxygen exposure inibits the polymerization process. However, only the edges of your gel will be affected by air exposure, and the middle of the gel will polymerize well.)

Step 6. Expand the Gel: Pry apart the two slides using foreceps, a razor blade, or some other thin edge. Put ~ 0.5 cm of deionized water in a petri dish or other container. Wet the paintbrush with water, use it to gently peel the gel off the slide and transfer it into the container with water. Let it sit in the water for ~10 mins, then remove the water and replace it with fresh water. Repeat this wash step 2-3 more times. The gel will expand ~4-fold. Be careful not to suck the gel up into a pipettor while changing the water!

Step 7. Play around with the gel to see how to pick it up and transfer it to other containers. The gels are very delicate when expanded, so this takes some finesse. Try picking up the gel by pushing it onto a coverslip using a paintbrush. If the gel is too large for this, cut it to make it smaller.

Exercise 2: Expansion of antibody-stained E. coli

Reagents (in addition to those used in Exercise 1):

Note 1: Acryoyl-X is an NHS ester, and is therefore very sensitive to water. It should be dissolved in anhydrous DMSO at 10 mg/mL and stored in 1-5 ul aliquots in PCR tubes in a desiccated container (eg bag or large tube with desiccation packets) at -20 degrees C.

Step 1. Grow E. coli overnight in liquid culture.

Step 2. The next day, pre-chill a solution of 70% ethanol, 1X PBS at -20 degrees C. (Eg for 1 mL, 100 ul 10X PBS, 700 ul ethanol, 200 ul pure water)

Step 3. Take 1.5 mL of mid log-phase culture and spin it down to form a pellet. This can be done using, for example, by spinning down at 7500xg for 10 mins. If possible, keep the cells cold at 4 degrees C during this step. Depending how long your cells were growing, you may need to use more of your culture to form a pellet. An ideal pellet is ~ 40 ul in volume.

Step 4. Remove supernatant and wash the cells by resuspending in PBS and then pelleting again. If possible, keep the cells cold at 4 degrees C during this step.

Step 5. Fix the cells by resuspending in pre-chilled 70% ethanol, 1x PBS and keeping the cells in fixation solution in the freezer for 15 minutes at -20 degrees.

Step 6. Spin down the cells, remove the supernatant, and resuspend in PBS. Repeat this two more times, so the cells are washed 3 times.

Step 7. Permeabilize the cells by resuspending in 1% Triton X-100, 1x PBS for 10 mins at room temperature. Then wash the cells once by pelleting, resuspending in PBS, and pelleting again.

Step 8. Set aside half the cells to be expanded later without antibody staining and used in Exercise 3, and move to step 9 using the other half.

Step 9. Block the cells by resuspending in 1% bovine serum albumin, 1X PBS for 15 mins. Then wash once in PBS.

Step 10. Stain the cells with primary anti-E. coli antibody by incubating in a 1:50 dilution of Abcam ab137967 in 1x PBS, 1% bovine serum albumin overnight at 4 degrees C. Depending on the size of your pellet, you may want to discard some of the pellet to reduce the amount of antibody you use. An acceptable final volume of antibody staining solution is ~200 ul, with a ~ 10 ul pellet suspended in it.

Step 11. Wash the cells 3 times in 1x PBS.

Step 12. Stain the cells with secondary goat anti-rabbit antibody by incubating in a 1:200 dilution of Abcam ab150077 in 1x PBS, 1% bovine serum albumin for 4 hours at room temperature.

Step 13. Wash the cells twice in 1x PBS.

Step 14. Add gel-linkable moieties to the antibodies on the cells by incubating the cells in 0.1 mg/mL acryloyl-X in PBS at room temperature for at least 6 hours, up to overnight. (0.1 mg/mL acryloyl-X can be made by diluting stock acryloyl-X (10 mg/mL in DMSO) 1:100 in PBS.)

Step 15. Wash the cells twice in PBS. During pelleting after the second wash, also pellet the cells that were set aside in step 8. Remove the supernatant from both tubes of cells and set the pellets aside to be used in step 17.

Step 16. Prepare for casting expansion gels by prepare two gelation chambers as described in step 3 of exercise 1, and place gel monomer solution (prepared in step 1 of exercise 1) and 10% APS and 10% TEMED on ice.

Step 17. (This step should be done as quickly as possible to avoid premature polymerization of the gel.) On ice, add 1 ul each of 10% TEMED and 10% APS to 48 ul of monomer solution and mix immediately by vortexing or aspirating repeatedly with the pipettor. Then resuspend the antibody-stained pellet in the gel solution you just made. Then use this solution to cast a gel as in step 5 in exercise 1. Repeat this also for the unstained cells, which will be used in exercise 3.

Step 18. Digest the cells by incubating the gels in Proteinase K diluted 1:100 (8 units/mL) in digestion buffer (recipe below) overnight at room temperature or at 37 degrees C for 4 hours. This can be done in a 30 mm petri dish or other small container with a flat bottom, or in a microfuge tube. (If you plan to image using a conventional fluorescent microscope, a 6-well glass-bottom dishes is an excellent container in which to digest and expand gels.) The volume of digestion buffer should be at least 100-fold the volume of the gel. If your gel is too large for this, cut it with a razor or other sharp edge to make it smaller. Gels can be transferred to digestion buffer using a paintbrush, similar to step 6 in exercise 1.

Digestion buffer:

Step 19. Expand the cells by washing the gels in deionized water 4 times for 10 minutes each, as in step 6 from exercise 1. This can be done in a petri dish or any container with a flat bottom. Note that the width of the chamber should be at least 4 times as wide as the original gel so there is room for it to expand. If your gel seems too large for this, you may wish to cut it to make it smaller. Also note that if you are transferring the gel from one vessel to another, you should be very gentle with it. The gels are delicate. They can be picked up most easily by pushing them onto a coverslip using a paintbrush.

Step 20. Transfer the gel onto a thin glass surface that will be used for imaging. (The gel can be picked up most easily by pushing it onto a coverslip, as mentioned above.) If using the mini microscope, this can be a coverslip. If you'll be using a conventional fluorescent microscope, use a coverslip or any other vessel with coverslip-thickness glass on the bottom that can be mounted on your microscope for imaging.

Step 21. Image the sample using fluorescence microscopy. Alexa 488, the fluorophore on the secondary antibodies that were retained in the gel, has peak excitation at 490 nm and peak emission at 525 nm, so it can be imaged using a standard 488 imaging channel. (This is the same imaging channel used to image GFP.) A lens with 40x or greater magnification is most ideal. The bacterial cells may be distributed all throughout the thickness of the gel, or they may have settled to the bottom of the gel, depending how quickly the gel polymerized relative to the sedimentation rate of the bacteria. If you can't see any bacteria in your gel, try flipping the gel over; the cells may be on the side of the gel furthest from the objective, which may be too far from the objective to see depending on its working distance.

Fluorescent In Situ Hybridization of expanded bacterial genomes

Prepare the FISH probes using nick translation

Background: Jena Bioscience Atto488 Nick Translation Labeling Kit contains all reagents (except template and materials for purification of the probe) required for nick translation labeling providing a highly efficient, easy-to-perform and rapid labeling technology.

The kit is recommended for direct enzymatic labeling of DNA. The Atto488 NT labeling mix contains specially optimized Atto488-XX-dUTP for incorporation into DNA by nick translation using DNA Polymerase I. The excellent stability and quantum yield of the fluorophore combined with a high incorporation rate of the dye-dUTP complex makes it the ideal choice for a broad range of fluorescence applications.

Nick translation labeling is based on the reverse activities of Polymerase I and DNase I. DNase I is able to introduce randomly distributed nicks to double stranded DNA at low enzyme concentrations. The 5'→3' exonuclease activitiy of Polymerase I removes nucleotides from the 3' side of the nick while synthesizing a partial new complementary strand using the 3'-OH termini as primer. In presence of dye-labeled dUTP Polymerase I incorporates labeled dUTP instead of dTTP. The well balanced polymerase / nuclease activities of the enzyme mix ensure the generation of highly labeled double stranded DNA fragments.

The resultant DNA is suited for application in FISH, microarray gene expression profiling and other nucleic acid hybridization assays.

Protect fluorescent labeled dUTP from light and carry out experimental procedures in low light conditions.

Kit Contents:

Enzyme mix (red cap): 2 units/μl polymerase I, 0.02 units/μl Dnase I in storage buffer

NT labeling buffer (green cap): 10x conc.

Atto488 NT labeling mix (purple cap): 0.5 mM dATP, 0.5 mM dCTP, 0.5 mM dGTP, 0.25 mM dTTP, 0.25 mM Atto488-XX-dUTP, pH 7.5

Stop buffer (yellow cap): 0.5 M EDTA, pH 8.0

PCR-grade water (white cap)

Prepare FISH Probes

1. Isolate genomic e. coli DNA.

Follow the instructions in the Qiagen QIAamp DNA Mini Kit or similar DNA clean-up kit or online DIY protocol, depending on resources. There are many ways to prepare purified genomic DNA, including phenol-chloroform isolation, silica columns, and more. All you need is ~1 ug of clean DNA for the nick translation kit. The E.coli chromosome is just over 4.5 MB in size, amounting to approximately .005 picograms per cell. A typical overnight culture from a single starting colony will contain approximately 1-2×10^9 cells/ml. Theoretically, that means that 1 ml of culture should yield about 5 µg of gDNA per 10^9 bacterial cells.

The input material for the nick translation reaction can be quantified by UV absorbance or estimated by gel electrophoresis (by comparing band intensity to a ladder of known concentration).

1. Prepare the Reaction Mix:

• Vortex the mix gently to assure homogeneity and centrifuge briefly to collect the reaction mixture at the bottom of the tube.
• Place the tube in a precooled thermomixer at 15 °C. An incubation of 90 min is recommended to generate DNA fragments in a size range between 200 and 500 bp.
• To control the length of the fragments load 2 μl of the assay on an agarose gel. Place the reaction tube at -20 °C while running the gel.
• To get smaller fragments add additional 2 μl of the Enzyme mix and extend the incubation at 15 °C.
• For final stopping the reaction add 5 μl of Stop buffer (yellow cap). Proceed to purification or store at -20 °C.
• Labeled DNA (1/10 of reaction) can be used for hybridization without purification.
• Optimal hybridization results are obtained when probe fragments are between 150 and 500 bp. You can use the methods from the Next Generation Synthesis class to determine the size of your FISH probes.

FISH Experiment

Use an expanded bacterial sample from the previous step, but without antibody staining (the antibody and the FISH probe use the same fluorescent color).

1. Denaturation of the probe

a) Denature the probe by heating the mix for 5 minutes at 73 ̊C in a water bath or a heating block with heated lid. b) Place the tube on ice for ~ 2 min. Centrifuge briefly. c) Pre-warm the denatured probe mix at 37 ̊C for 15 minutes before hybridization.

1. Hybridization of the probe

a) Bring sample to room temperature. b) Pre-warm the denaturation buffer (70% formamide, 2x SSC, pH 7.0-8.0) at 75 ̊C for 30 min. Immerse the sample into enough denaturation solution to completely submerge the gel, incubate for 5 minutes. Remove excess denaturation buffer. c) Submerge the gel in hybridization buffer (10% dextran sulfate, 30% formamide, 2X SSC) and add the denatured probe mix (from step 1c). Incubate overnight to 72 hours at 42 ̊C.

1. Post Hybridization Wash

a) Pre-warm Wash Solution (0.4X SSC, 0.3% Triton-X 100) at 73 ̊C for 30 minutes. b) Submerge gel in Wash Solution for 30 minutes at room temperature. b) Expand gels by submerging in 0.05X SSC. You may need to exchange the expansion buffer several times to achieve full expansion & reduce background signal.

Image the gel using the mini-microscope to visualize the individual bacterial cells labeled by fluorescent genomic DNA FISH.

Resources