TITLE: AAV-Zombie on tissue sections
AUTHORS: Gerard Michael Coughlin

[DESCRIPTION]
Detection of AAV genomes in situ can facilitate understanding of AAV transduction and processing. This protocol enables spatial detection of AAV genomes and concatemers in tissue and is based on the Zombie method published in Askary et al., Nat Biotechnol 38: 66. AAVs carrying barcoded genomes are delivered to cells. Following fixation, a phage RNA polymerase is used to transcribe the barcode in situ, yielding barcoded transcripts that can be detected with RNA FISH. This protocol is compatible with either ssAAV or scAAV genomes. In the former case, detected genomes will be primarily nuclear and detectable once second strand synthesis has occurred. In the latter case, the genome can be detected regardless of where in the transduction pathway the genome is. This protocol can be combined with RNA FISH detection of reporter transcripts or endogenous transcripts, enabling measurement of expression and cell type classification.

[BEFORE_START]
This protocol enables spatial detection of AAV genomes and concatemers in tissue. For detection of AAV genomes, a Zombie barcode and phage RNA polymerase promoter are required in a single AAV genome. For detection of AAV concatemers (SpECTr), 2 AAV genomes are required: one genome carries a Zombie barcode and the other carries a phage RNA polymerase promoter. Barcode sequences can be found in Supplementary Table 1 of Askary et al., Nat Biotechnol 38: 66.

See https://www.biorxiv.org/content/10.1101/2023.12.23.573214v1 for more information on these methods. 

[STEPS]
SECTION: Reagent set up
1. General note on reagents and consumables

 

SECTION: Sample preparation
15. On day of tissue harvest, prepare work area. In addition to tools and equipment for transcardial perfusion and tissue dissection, you will need:

ice-cold 1x PBS
clean 10 cm petri dishes
ice
tissue embedding molds
O.C.T. compound
dry ice
dry ice-ethanol slurry

Before beginning tissue collection, label all tubes and embedding molds with solvent-resistant marker. Place a small amount of O.C.T. compound into the bottom of each embedding mold. 

If analysis of fixed tissue is also desired (e.g. for detection of an AAV-encoded fluorescent protein), prepare tubes with appropriate volumes of 4% PFA in 1x PBS and keep on ice. In addition, tissue may be collected and homogenized, dissociated, or snap-frozen for other downstream analyses.     

 

SECTION: Sample preparation
14. Transduce animals with AAVs carrying Zombie barcode and phage RNA polymerase promoter (barcode sequences can be found in Supplementary Table 1 of Askary et al., Nat Biotechnol 38: 66).

 

SECTION: In situ transcription and probe hybridization
22. General note on section

 

SECTION: Reagent set up
2. Prepare 3:1 methanol:acetic acid (MAA) fixation buffer by combining:

Methanol: 180 mL 
Glacial acetic acid: 60 mL 

in a 250 mL sterile bottle. Store solution at -20 °C  for up to 3 months.

 

SECTION: Reagent set up
3. Prepare 70% ethanol solution, by combining

100% Ethanol: 175 mL 
UltraPure water: 75 mL 

in a 250 mL sterile bottle. Store solution at -20 °C for up to 3 months.

 

SECTION: Reagent set up
5. Prepare probe hybridization buffer, by combining:

20x saline-sodium citrate (SSC) buffer: 5 mL 
Ethylene carbonate: 5 mL 
50% dextran sulfate: 10 mL 
UltraPure water: 30 mL 

in a 50 mL conical tube. Store solution at Room temperature for up to 3 weeks, or -20 °C for up to a year. 

 

SECTION: Reagent set up
6. Prepare stringent wash buffer, by combining:

20x saline-sodium citrate (SSC) buffer: 5 mL 
Ethylene carbonate: 15 mL 
UltraPure water: 30 mL 

in a 50 mL conical tube. Store solution at Room temperature for up to 3 weeks, or -20 °C for up to a year. 

 

SECTION: Reagent set up
4. Prepare 4% paraformaldehyde, 1x PBS solution (4% PFA, 1x PBS), by combining:

32% paraformaldehyde: 25 mL 
10x Phosphate-buffered saline (PBS): 20 mL 
UltraPure water: 155 mL 

in a 250 mL sterile bottle. Store solution at 4 °C for up to 1 month. 
SECTION: Reagent set up
7. Prepare wash buffer (5x SSC, with 0.1% Tween-20), by combining:

20x saline-sodium citrate (SSC) buffer: 100 mL 
10% Tween-20: 5 mL 
UltraPure water: 395 mL 

Store solution at Room temperature for up to a year.

SECTION: Reagent set up
9. Prepare HCR amplification buffer, by combining:

20x saline-sodium citrate (SSC) buffer: 5 mL 
Ethylene carbonate: 5 mL 
UltraPure water: 35 mL 

in a 50 mL conical tube. Store solution at Room temperature for up to 3 months.

 

SECTION: Reagent set up
10. Prepare 1x PBS, by adding:

10x PBS: 55.5 mL 

To a 500 mL bottle of UltraPure water. 

SECTION: Sample preparation
13. General note on sample preparation and scaling

 

SECTION: In situ transcription and probe hybridization
23. Prepare the staining tray for the in situ transcription reaction. Pour 100 mL of MilliQ (or comparable) water in the base of the tray to provide a humidified environment. Place the staining tray in 37 °C incubator for at least 60 min .

 

SECTION: In situ transcription and probe hybridization
24. Once staining tray is prepared, thaw the NTP(s) and buffer at Room temperature , then place on ice.  

 

SECTION: In situ transcription and probe hybridization
25. Set up in situ transcription reaction solution. For each slide, prepare 200 µL of reaction solution. If using T7 RNA polymerase, prepare the following reaction mix:

NTPs: 80 µL (or 20 µL of each NTP, if storing separately)
UltraPure water: 80 µL 
RNA polymerase: 20 µL 
Buffer: 20 µL 

If using SP6 RNA polymerase, prepare the following reaction mix:

NTPs: 80 µL (or 20 µL of each NTP, if storing separately)
UltraPure water: 60 µL 
RNA polymerase: 40 µL 
Buffer: 20 µL

Store reaction solution on ice until use.

 

SECTION: In situ transcription and probe hybridization
27. Retrieve samples from -20 °C storage. Decant the 70% ethanol into a waste beaker, then fill the jar with 1x PBS. Decant wash then refill jar with 1x PBS and keep at Room temperature until use. 
SECTION: In situ transcription and probe hybridization
26. Set up tools and disposables. You will need:

Kimwipes
A pair of clean forceps (suitable for lifting and manipulating slides)
Appropriately sized coverslips (for a 25 mm x 75 mm slide full of tissue sections, use a 24 mm x 60 mm coverslip)
A 50 mL tube with 50 mL of UltraPure water. 
SECTION: In situ transcription and probe hybridization
28. Start in situ transcription reactions.

Remove staining tray chamber from 37 °C incubator. 
Lift each slide out of Caplin jar, wash briefly by submerging in UltraPure water, then remove excess water from slide by leaning slide against a vertical surface for 30 s 
Gently tap bottom of slide against Kimwipe or a piece of paper towel on the bench surface to remove  pooled water, then dry the back of the slide with a Kimwipe. Place slide onto the staining tray, with the tissue facing up.
Pipette 200 µL of reaction mixture (Step 24) onto the slide
Spread the reaction mixture over the tissue by gently lowering a clean coverslip over the tissue, being careful not to introduce bubbles
Repeat for each slide

SECTION: In situ transcription and probe hybridization
29. Transfer staining tray to 37 °C incubator, and incubate for 180 min .

 

SECTION: In situ transcription and probe hybridization
30. During in situ transcription, pre-warm the hybridization buffer to 42 °C . You will need 1.5 mL of hybridization buffer per slide.

 

SECTION: In situ transcription and probe hybridization
31. Once the in situ transcription is complete, fix nascent transcripts in place with PFA. Fill a Coplin jar with ice-cold 4% PFA in 1x PBS. Retrieve staining tray from incubator. Pick up each slide and hold vertically, allowing the coverslip to slide off. 

Transfer to the 4% PFA, 1x PBS, and fix for 30 min at Room temperature .

 

SECTION: In situ transcription and probe hybridization
32. Following fixation, transfer slides to a new Coplin jar filled with 1x PBS. Wash twice in 1x PBS to remove residual PFA, then wash twice in 5x SSC (5 min per wash).

Lift each slide out of Caplin jar and remove excess buffer by leaning slide against a vertical surface for 30 s . Gently tap bottom of slide against Kimwipe or a piece of paper towel on the bench surface, then dry the back of the slide with a Kimwipe. Place slide onto the staining tray, with the tissue facing up.

Add 1 mL of pre-warmed hybridization buffer, ensuring that all the tissue is covered by hybridization buffer. Incubate at 37 °C for 60 min . 
SECTION: In situ transcription and probe hybridization
33. During incubation in Step 31, prepare probe solution by diluting probes to 4 nM  in the remaining hybridization buffer. Return to 42 °C until use. 

 

SECTION: In situ transcription and probe hybridization
34. Following equilibration in hybridization buffer, remove old hybridization buffer from slides by tilting over paper towel. Remove excess hybridization buffer by leaning slide against a vertical surface for 30 s . Gently tap bottom of slide against Kimwipe or a piece of paper towel on the bench surface to remove pooled hybridization buffer. 

Return slides to staining tray, and add 500 µL of prewarmed probe solution (Step 32). Spread the probe solution over the tissue by gently lowering a clean coverslip over the tissue, being careful not to introduce bubbles.

Incubate at 37 °C   .

 

SECTION: Probe washes and Hybridization chain reaction (HCR) amplification
35. Note on reaction volumes

 

SECTION: Probe washes and Hybridization chain reaction (HCR) amplification
36. Pre-warm stringent wash buffer to 37 °C , vortexing a couple times to ensure components are thoroughly mixed. You will need 3 mL of stringent wash buffer per slide. 
SECTION: Reagent set up
8. Prepare 5x SSC, by combining:

20x saline-sodium citrate (SSC) buffer: 100 mL 
UltraPure water: 400 mL 

Store solution at Room temperature for up to a year

SECTION: Probe washes and Hybridization chain reaction (HCR) amplification
37. Once stringent wash buffer is warmed to 37 °C and mixed, begin stringent washes. 

Retrieve staining tray from incubator. Remove coverslip by holding slide vertically over a piece of paper towel, allowing the coverslip to slide off. Remove excess probe solution by leaning slide against a vertical surface for 30 s . Gently tap bottom of slide against Kimwipe or a piece of paper towel on the bench surface to remove pooled hybridization buffer. 

Return slides to staining tray and pipette 1 mL of stringent wash buffer onto the slide. After 1 min, remove that rinse by gently lifting slide and dumping stringent wash buffer into a waste collection beaker. Return slide to staining tray, then place staining tray into the 37 °C  incubator, then add 1 mL of stringent wash buffer. Place lid on staining tray and incubate for 30 min. 

Repeat wash step 1 more time.

 

SECTION: Probe washes and Hybridization chain reaction (HCR) amplification
38. Retrieve staining tray from incubator. Remove stringent wash buffer from slides by decanting slide into waste beaker, then lean slide against a vertical surface for 30 s . Gently tap bottom of slide against Kimwipe or a piece of paper towel on the bench surface to remove pooled stringent wash buffer. 

Transfer slides to a Coplin jar. Wash slides in wash buffer (5x SSC, with 0.1% Tween-20). Perform 1 brief rinse in wash buffer, followed by two longer washes at 37 Room temperature for 30 min each. 
SECTION: Probe washes and Hybridization chain reaction (HCR) amplification
39. Following washes, remove slides from Coplin jar and lean against a vertical surface for 30 s . Gently tap bottom of slide against Kimwipe or a piece of paper towel on the bench surface to remove pooled wash buffer. 

Dry the back of the slides with a Kimwipe, then return to staining tray. Add 1 mL of HCR amplification buffer, ensuring that the buffer distributes over the slide surface. Equilibrate samples in HCR amplification buffer for   at Room temperature .
SECTION: Probe washes and Hybridization chain reaction (HCR) amplification
40. During HCR amplification buffer equilibration step, prepare HCR hairpins.

Aliquot each hairpin separately into PCR tubes
Using a thermocycler, heat hairpins to 95 °C for 2 min 
Remove hairpins from thermocycler and allow to cool back down to room temperature in the dark (at least 10 min 

 

SECTION: Probe washes and Hybridization chain reaction (HCR) amplification
41. Following equilibration in HCR amplification buffer, prepare HCR amplification solution. For each slide, prepare 300 µL of amplification solution. Dilute snap-cooled hairpins to 60 nM in the HCR amplification buffer. 

For example, for one coverslip with one probe set using the B5 initiator, combine:

HCR amplification buffer: 288 µL 
Hairpin 1 (e.g. B5H1): 6 µL 
Hairpin 2 (e.g. B5H2): 6 µL 

Briefly vortex to mix. 

 

SECTION: Probe washes and Hybridization chain reaction (HCR) amplification
42. Remove old amplification buffer from slides by tilting over a piece of paper towel or waste collection beaker. Return slides to staining tray and add 300 µL of amplification solution. Spread the HCR amplification solution over the tissue by gently lowering a clean coverslip over the tissue, being careful not to introduce bubbles. Incubate 60 min to 180 min at Room temperature and in the dark.

 

SECTION: Probe washes and Hybridization chain reaction (HCR) amplification
43. Following HCR amplification, remove coverslips by holding the slide vertically, allowing the coverslip to slide off. 

Remove unassembled hairpins by washing 4 times slides in wash buffer (5x SSC, with 0.1% Tween-20), in a Coplin jar. Perform 1 brief rinse with wash buffer, followed by three longer washes at 37 Room temperature for 10 min each. 
SECTION: Probe washes and Hybridization chain reaction (HCR) amplification
45. (Optional) Label nucleus with Hoescht 33342. 

For each slide, prepare 1 mL of 1x PBS with 1/10000 Hoechst 33342. Rinse slides briefly in 1x PBS in a Coplin jar, then return to staining tray. Pipette 1 mL  of Hoechst solution directly onto slide and incubate for 30 min at Room temperature in the dark. 

Following incubation, wash samples 3 times with 1 mL of 1x PBS at 37 Room temperature for 5 min each. 

 

SECTION: Probe washes and Hybridization chain reaction (HCR) amplification
46. Mount coverslips onto slides. You will need a 50 mL tube with 50 mL of UltraPure water, a suitable mounting medium, and appropriately-sized glass coverslips.

For each slide:
Remove slide from Coplin jar, and immerse briefly in UltraPure water
Transfer slides to a fume hood, allowing the tissue to dry to the point of being transparent
Once tissue is dry, pipette 100 µL of mounting media in a line on the surface of the slide, near the tissue sections
Spread the mounting media over the tissue by gently lowering a clean coverslip over the tissue, being careful not to introduce bubbles. 

Once all coverslips are mounted, gently press down on each coverslip to push out excess mounting media. Store in the dark at Room temperature for at least    until mounting media cures. Once cured, excess Prolong Diamond can be easily removed by running slides under a gentle stream of MilliQ water, being careful to not displace coverslips. 

Samples are now ready to image. 

 


SECTION: Probe washes and Hybridization chain reaction (HCR) amplification
44. (Optional) Label proteins with immunofluorescence staining. 

We expect that most standard immunofluorescence protocols can be used here. Below is the protocol we used for labeling AAV capsid proteins with immunofluorescence using a fluorophore-conjugated primary antibody. Optimal incubation conditions, dilutions, and durations, will need to be determined empirically for each protocol and antibody.

Block slides in 1 mL of BlockAid blocking buffer with 0.1% Triton X-100, for 60 min at Room temperature 
Incubate samples with 1:100 dilution of primary antibody in BlockAid blocking buffer with 0.1% Triton X-100, 4 °C for   .
Wash samples 3 times in 1x PBS-Tx, for 15 min each at Room temperature 

 

SECTION: Reagent set up
12. (Optional) Prepare 1x PBS-Tx, by combining:

10x PBS: 50 mL 
10% Triton X-100: 5 mL 
UltraPure water: 445 mL 

 

SECTION: Sample preparation
16. Begin tissue collection.  Euthanize animals according to your lab's approved protocol, then transcardially perfuse animals with 30 mL of heparinized PBS. Working quickly, dissect out and place tissue of interest into a petri dish with ice-cold PBS, then isolate regions of interest. For AAV-Zombie, dissect tissue into small blocks (max size 0.5 cm x 0.5 cm x 0.5 cm), then place into embedding mold. Using a clean pipette tip, gently position tissue. Once the mold is full, add enough O.C.T. compound to cover the tissue, then lower the mold into the dry ice-ethanol slurry. Be careful not to let any ethanol spill into the O.C.T. Once the entire block is frozen, transfer the block to dry ice (or to a -70 °C freezer). 

If also collecting tissue to fix, gently drop tissue into 4% PFA in 1x PBS, and post-fix at 4 °C , for   , then proceed with standard procedures. Fresh tissue may also be harvested and homogenized, dissociated or snap-frozen for other downstream analyses.

 

SECTION: Sample preparation
17. Slice tissue. Remove flash frozen blocks of tissue from -70 °C , and allow to warm to cryostat temperature. Section tissue at 20 µm  and collect sections on clean glass slides. Allow sections to dry on the top of the cryostat, then transfer to a slide box, seal with tape, and place into -70 °C storage until use. 

 

SECTION: Tissue MAA fixation
18. Remove slide box from -70 °C storage and allow to warm to room temperature before removing tape and opening. 

 

SECTION: Tissue MAA fixation
19. Briefly wash slides by submerging them in 1x PBS, in a Coplin jar, for 5 min . This step is to remove O.C.T. from the slide before fixation. Leave slides in 1x PBS until ready for fixation step. 

SECTION: Tissue MAA fixation
20. Fix sections in MAA. Fill a Coplin jar with ice cold MAA fixation solution. Remove slides from 1x PBS, and hold vertically to allow excess buffer to run off the slide. Touch slide to a piece of paper towel to further remove excess buffer. Submerge slides in MAA, and transfer to -20 °C freezer to fix for 180 min .

 

SECTION: Tissue MAA fixation
21. Following fixation, decant MAA into an appropriate container for disposal. Fill Coplin jar with ice-cold 70% ethanol. Decant wash ethanol into MAA waste container, and fill Coplin jar again with ice-cold 70% ethanol. Transfer samples to -20 °C until processing. 

 

SECTION: Reagent set up
11. Prepare heparinized PBS, by combining:

1 PBS packet
Heparin sodium salt: 50 mg 
MilliQ water (or equivalent): 1 L 

Shake to combine and store at Room temperature for up to 3 months.