Protocol Tag: HBV Nucleic Acid Analyses

Sample preparation

1. Cell culture and HBV infection.

1.1 Tetracycline-inducible (tet-off) HepAD38 were seeded on collagen-coated four-well chamber slides and maintained in culture medium for 48 – 72 h.

1.2 HepG2-NTCP cells were cultured in collagen-coated four-well chamber slides, and at approximately 100% confluence, concentrated supernatants of HepAD38 cells were mixed with culture medium containing 4% PEG8000 and inoculated onto cells for 6 – 8 h. The cells were subsequently washed three times with PBS and maintained in replication medium for several days based on the aim of experiment.

2. Fix the cells in 3.7% formaldehyde solution and dehydration.

2.1 In a fume hood, prepare 6 ml of fresh 3.7% formaldehyde solution by diluting 500 ul of a 37% stock formaldehyde with 5.5 ml of PBS and then mixing well.

▲Caution: Formaldehyde is a poison and irritant. Avoid contact with skin and mucous membranes.

2.2 Gently aspirate off culture medium and rinse cells three times, each time with 300 ul/well of PBS.

• Critical: Before fixation, cell attachment may be weak. Aspiration and dispensing of contents from and into the chamber should be performed slowly and gently with a 1 mL pipette.

Aspirate from the corners and dispense against the chamber wall.

2.3 Aspirate off the final PBS wash and add 300 μL/well of freshly prepared 3.7% formaldehyde solution. Cover chamber slide with lid and incubate at RT for 10 minutes.

2.4 Aspirate off the formaldehyde solution and gently rinse the cells three times, each with 300 ul/well of PBS.

2.5 Aspirate off the PBS and replace it with 300 μL/well of 50% ethanol. Incubate for 5 minutes at RT.

2.6 Aspirate off the 50% ethanol and replace with 300 μL/well of 100% fresh ethanol. Seal the chamber slides with parafilm and store the dehydrated cells in 100% ethanol at –20 °C until used^a.

If doing the experiment right away, aspirate off the 50% ethanol and replace with 300 μL/well of 70% ethanol at 4 °C for at least 1 hour and proceed on 3.1.B

NOTE:

1. Dehydrated cells can be stored under these conditions for up to one month and must be rehydrated before being used in the in situ assay.

FISH assay procedure

3. Cell rehydration.

3.1 A. Aspirate off the 100% ethanol (corresponding to 2.6 A) and replace with 300 μL/well of 70% ethanol. Incubate for 5 minutes at RT.

B.  Aspirate off the 70% ethanol (corresponding to 2.6 B) and replace with 300 μL/well of 50% ethanol. Incubate for 5 minutes at RT.

C. Aspirate off the 50% ethanol and replace with 300 μL/well of PBS. Incubate for 10 minutes at RT.

3.2 Aspirate off the PBS and add 300 μL/well of a 3.7% formaldehyde solution. Incubate for 10 minutes at RT.

3.3 Aspirate off the formaldehyde solution and gently rinse the cells three times, each with 300 μl/well of PBS for 5 minutes at RT.

4. Protease digestion^a.

4.1 Prepare 1.2 ml diluted working Protease Solution (1:3000) in PBS by adding 0.4 ul Protease QS^b to 1199.6 μl PBS. Vortex briefly to mix.

4.2 Aspirate off the PBS and added 300 μL/well of working protease solution. Cover chamber slides with lid and incubate for 10 minutes at 37 °C in a water bath.

4.3 Aspirate off the working protease solution from the chamber and rinse the cells three times, each with 300 μl /well of PBS.

4.4 Aspirate off PBS and add 300 μL/well of a 3.7% formaldehyde solution. Incubate for 5 minutes at RT.

4.5 Aspirate off the formaldehyde solution and gently rinse the cells three times, each with 300 μL/well of PBS.

NOTES:

1. The Protease digestion step partially degrades capsid and polymerase that wrap HBV DNA thus increasing target accessibility. However the exact concentration of protease should be empirically determined because excessive digestion can cause complete release of nucleic acids and even cell morphology damage and cell loss.
2. Protease QS is provided in the kit. Thaw and place Protease QS on ice until use.

 

5. Digest with RNase A/H^a.

5.1 Prepare 1.2 ml diluted working RNase A/H solution by adding 2.4 μl RNase A 1:500 and 2.4 μL RNase H 1:500 to 120  μL 10X RNaseH Reaction buffer and 1075.2 μl deionized water.

5.2 Aspirate off the PBS and added 300 μL/well of working RNase A/H solution. Incubate the slides in a cassette for 1 hour at 37 °C on the water bath.

5.3 Aspirate off the working RNase A/H solution and gently rinse the cells three times, each with 300 μL/well of PBS for 5 minutes at RT.

5.4 Aspirate off the PBS and added 300 μL/well of 3.7% formaldehyde solution. Incubate for 5 minutes at RT.

5.5 Aspirate off the formaldehyde solution and gently rinse the cells three times, each with 300 μL/well of PBS for 5 minutes at RT.

NOTES:

1. a. RNase A is an endonuclease that specifically degrades single-stranded RNA.

RNase H is an endonuclease, it can decompose the RNA strand in the RNA/DNA hybrids.

Our previous results suggested that HBV RNA could exist in the form of RNA/DNA hybrid in core particles^1,3.  The combined use of RNase A/H digestion can increase signal and improve assay specificity. First, probe set 3 targets the plus strand of HBV DNA but can also bind to HBV RNA. RNase digestion eliminate residual HBV RNA in the replication complex and capsid-free HBV mRNA. Second, RNaseH digestion eliminate the HBV RNA that binds to minus strand DNA hence increasing the hybridization efficiency of probe set 2.

 

   6. Hybridization with Probe.

6.1 Prepare 150 μl  working Probe Set Solution by diluting 1.5 μl of each Probe Set (1:100) in 148.5 μl  pre-warmed Probe Set Diluent QF^a. Gently mix the solution and shortly centrifuge 10-20 seconds. 150 µl for every slide?

▲Caution: Probe Set Diluent QF contains formamide, a teratogen, irritant and possible carcinogen. Avoid contact with skin and mucous membranes.

6.2 Remove the slides from the chamber and gently decant the remaining PBS, then put the slide edge on a clean, dry paper towel for 1-2 seconds. Add 150 μl/slide of the working Probe Set Solution to the slide, subsequently cover with a 24×50 mm² coverslip and seal edges with rubber cement.

• Critical: The chamber on the slide may not be easily removed. Handle with care to avoid the slide broken.

6.3 Preheat the cassette at 75 °C in the dry incubator for 2 minutes.

6.4 Then incubate at 40 °C for 3 hours in the water bath or the Thermobrite hybridizer.

NOTE:

1. Pre-warm Probe Set Diluent QF to 40 °C in a water bath for 10 minutes to dissolve possible precipitates.

 

7. Hybridize with Pre-Amplifier

7.1 Prepare 15 ml Wash Buffer by adding 45 μl Wash Comp1 and 75 μl Wash Comp2^a to 14.88 ml deionized water and then mixing well.

7.2 Prepare 150 μl  working Pre-Amplifier Mix Solution by diluting 0.75 μl Pre-Amplifier Mix 1:200 in 149.25 μl pre-warmed Amplifier Diluent QF. Gently mix the solution and transient centrifuge for 10-20 seconds.

▲Caution: Amplifier Diluent QF contains formamide, a teratogen, irritant and possible carcinogen. Avoid contact with skin and mucous membranes.

7.3 Remove rubber cement and coverslips from the slide, then put the slide edge on a clean, dry paper towel for 1–2 seconds. Wash slides three times, each with 1 mL/slide of Wash Buffer. Soak sample for 5 minutes during each wash.

• Critical: During all subsequent washing steps, the slides should be washed fully with wash buffer to eliminate non-specific binding, but do not soak samples in Wash Buffer longer than 30 minutes. Are all the washing steps at room temperature?

7.4 Gently remove the Wash Buffer, put the slide edge on a clean, dry paper towel for 1–2 seconds. Add 150 μl/slide of the working PreAmplifier Mix Solution to the slide, then cover with a 24×50 mm² coverslip.

7.5 Incubate at 40 °C for 35 minutes in the water bath.

NOTES:

1. Wash Comp 1 and Wash Comp 2 are provided in the kit, they should be free of precipitates before making a working wash buffer.

 

8. Hybridize with Amplifier

8.1 Prepare 150 μl  working Amplifier Mix Solution by diluting 6 μl Amplifier Mix 1:25 in 144 μl pre-warmed Amplifier Diluent QF. Gently mix the solution and transient centrifuge for 10-20 seconds.

▲Caution: Amplifier Diluent QF contains formamide, a teratogen, irritant and possible carcinogen. Avoid contact with skin and mucous membranes.

8.2 Remove coverslips from the slide, then put the slide edge on a clean, dry paper towel for 1–2 seconds. Wash slides three times, each with 1 mL/slide of Wash Buffer. Soak sample for 5 minutes during each wash.

8.3 Gently aspirate the Wash Buffer, put the slide edge on a clean, dry paper towel for 1–2 seconds. Add 150 μl /slide of the working Amplifier Mix Solution to the slide, then covered with a 24×50 mm² Cover Slip.

8.4 Incubate at 40 °C for 35 minutes in the water bath.

 

9. Hybridize with Label Probe

• Critical: Protect samples from light during all subsequent steps.

9.1 Prepare 150 μl  working Label Probe Mix Solution by diluting 6 μl  Label Probe Mix 1:25 in 144 μl  pre-warmed Label Probe Diluent QF. Gently mix the solution and shortly centrifuge for 10-20 seconds. Protect from direct light exposure.

9.2 Remove coverslips from the slide, then put the slide edge on a clean, dry paper towel for 1–2 seconds. Wash slides three times, each with 1 mL/slide of Wash Buffer. Soak sample for 5 minutes during each wash.

9.3 Gently aspirate the Wash Buffer, put the slide edge on a clean, dry paper towel for 1–2 seconds. Add 150 μl /slide of the working Label Probe Mix Solution to the slide, then cover with a 24×50 mm² coverslip.

9.4 Incubate at 40 °C for 25 minutes in the water bath.

 

10. Nuclear counterstaining

10.1 Prepare 500 μl  working Hoechst Solution by diluting the 0.5 μl  Hoechst 1:100 in 999.5 μl  PBS. Gently mix the solution. Protect from light.

Note: Nuclear staining can also be done with DAPI. Dilute the 100 X DAPI (provided in the ViewRNA ISH cell assay kit) 1:100 in nuclease free water to obtain a working DAPI solution and perform as follows.

▲Caution: DAPI is a possible mutagen. Avoid contact with skin and mucous membranes.

10.2 Remove coverslips from the slide, then put the slide edge on a clean, dry paper towel for

1–2 seconds. Wash slides three times, each with 1 mL/slide of Wash Buffer. Soak sample for 5 minutes during each wash.

10.3 Gently aspirate the Wash Buffer, put the slide edge on a clean, dry paper towel for 1–2 seconds. Add 500 μl/slide of the working Hoechst Solution to the slide and incubate for 3 minutes at RT.

10.4 Decant working Hoechst Solution from the slide. Wash slides three times with 1 mL/slide of PBS. Soak sample for 5 minutes during each wash.

10.5 Aspirate PBS and add two drops of fluorescent mounting medium onto the slide, and finally cover with a coverslip to preserve a better fluorescent signal.

10.6 Use a clean paper towel to soak up the excess mounting medium on the sides of the slide when necessary, and seal the slide edges with nail polish.

10.7 Allow the nail polish to dry for at least 15 minutes.

 

 

11. Imaging

A. Samples may be viewed under a fluorescent microscope immediately.

        B. The slide should be stored at 2–8°C protected from light. Fluorescent signals will be stable for up to one week when stored properly.

        C. For viewing signals, use appropriate filter sets. Here we can use filter set Cy3 (550 nm) to detect Probe Set type 1^a or use filter set Cy5 (650 nm) to detect Probe Set type 6^b.

NOTES:

a. Signal of LP1-550 is visible to unaided eyes under epi-fluorescent microscope, and appear as red dots.

b. Signal of LP6-650 is invisible to unaided eyes under epi-fluorescent microscope and has to be captured by a digital camera.

c. Due to the relative weak signal of FISH, a digital camera with high quantum efficiency is desirable. In addition, high sensitivity mode (e.g. HyD mode in Leica SP8) must be used when using a confocal microscope. The signal is also more prone to be bleached due to the high energy of confocal laser beam. We have also imaged the FISH results using STORM (Type 6 probe only) and STED with some success. Generally, these methods generate higher resolution images but the procedure is much more complex.

d. If one wishes to see colocalization between nucleic acid and protein, a routine immunofluorescence protocol can be performed immediately after 10.2 with no need for fixation or permeabilization step.

 

 

12. Typical results

Typical imaging results were shown below². Signal generated by probe set 2 (green) indicates all viral DNA (including core particle DNA and deproteinated DNA), signal generated by probe set 3 (red) indicates deproteinated rcDNA and some core particle DNA in the cytoplasm. Our previous results showed that the rcDNA in HepAD38 has longer plus strand which resulted in a diminished selectivity of probe set 3 toward intranuclear rcDNA and cccDNA. The yellow arrow indicates intranuclear HBV DNA that is detected by both probes. Further differentiation between episomal DNA and integrated HBV DNA is beyond the capability of the assay.

More images are also available at http://www.hepb-atlas.com/home/projects/detail/12

 

Cell Cultures

1. Seed HBV stable cell lines, such as HepG2.2.15 (5), or tetracycline-inducible (tet-off) HepAD38 (6), or tetracycline-inducible (tet-off) HepDE19/DES19 cells (7, 8), in collagen coated 6-well plates at approximately 100% confluence with appropriate culture medium.

Note: Collagen-coated plate is ideal for cells to form confluent monolayers, especially for HepG2-based cells.

Note: HepG2.2.15 cell line is cultured with DMEM/F12 medium supplemented with 10% fetal bovine serum, 100 U/mL penicillin and 100 μg/mL streptomycin, plus 400 μg/mL G418. Tetracycline-inducible (tet-off) cell lines are maintained in the same way as the HepG2.2.15, but with the addition of 1 μg/mL tetracycline.

2. When the cell monolayer reaches complete confluence (normally 24-48 hours post seeding), change the culture medium with fresh medium. For tetracycline-inducible cell lines, remove the tetracycline from culture medium to induce HBV replication.

Note: To obtain a high level of HBV replication in tetracycline-inducible cell lines, it is critical to withdraw the tetracycline when the cells reach confluence, as HBV replication will arrest cell growth.

Note: HBV infected cells can also be subjected to the following procedures.

 

cccDNA Extraction  from Cell Culture

1. Lyse cells by adding 1.5 mL of TE buffer (10:10) and 0.1 mL of 10% SDS into each well of a 6-well cell culture plate. Gently mix and incubate the plate for 30 min at room temperature.
2. Transfer the viscous cell lysate to a 15 mL centrifuge tube. Add 0.4 mL of 5 M NaCl and gently invert the tube 10 times. Let the tube sit at 4°C for at least 16 h to efficiently precipitate proteins and protein-associated DNAs.

Note: Use wide-mouth plastic transfer pipette to transfer the lysate into centrifuge tube.

3. Centrifuge at 14,500 × g for 30 min at 4°C. Transfer the supernatant to a fresh 15 mL centrifuge tube. To remove the residual protein from the supernatant, add an equal volume of phenol (approx. 2 mL) to the supernatant and mix thoroughly by hand shaking for 10 sec. Centrifuge at 3,500 × g for 10 min at 4°C, and transfer the aqueous phase to a fresh 15 mL tube. Repeat phenol extraction step. Add equal volume of phenol/chloroform (approx. 2 mL) and mix thoroughly by hand shaking, centrifuge at 3,500 × g for 10 min at 4°C and transfer the aqueous phase to a fresh 15 mL tube.

Note: Use leak-proof tube with screw cap. Do not shake the tube too vigorously or too long, which it may physically nick the supercoiled cccDNA.

4. Add two volumes of 100% ethanol (approx. 4 mL) and mix thoroughly by inverting the tube several times. Incubate at room temperature overnight to precipitate DNA.

Note: Do not incubate at low temperature. Because high concentration of salt is used in Hirt extraction, incubation of DNA solution at low temperature will precipitate high amount of salt.

5. On the third day, centrifuge the tube at 3,500× g for 30 min at 4°C, and discard the supernatant. Add an equal volume of 75% ethanol (approx. 2 mL) and gently flick the tube to wash the DNA pellet. Centrifuge at 3,500× g for 15 min at 4°C.
6. Discard the supernatant. Allow the pellet to air dry for about 10 min at room temperature. Dissolve the DNA pellet in 25 μL TE buffer (10:1).

 

Agarose Gel Electrophoresis

1. Prepare 150 mL of 1.2% gel by mixing 1.8 g agarose, 5 mL of 30× TAE buffer, and 145 mL of distilled water in a microwavable flask. Microwave the flask until the agarose is thoroughly dissolved. Let it cool down to about 50°C at room temperature. Pour the liquid gel into the gel tray (12cm×10cm) with an appropriate comb inserted. Let the gel solidify at room temperature before removal of the comb.
2. Prepare 600 mL of gel running buffer by diluting 20 mL of 30× TAE buffer with de-ionized water.
3. Prepare samples by adding approximately 3 μL of 10× loading buffer to 25 μL of each sample. Pipette up and down to mix well.
4. Assemble submarine gel running unit by correctly positioning the gel box, adding running buffer into the chamber, and loading 28 μL of each sample into a separate well. Connect the gel running unit to a power supply and run gel at 25 V for overnight.

 

Southern Blot

1. Disconnect the gel running unit and take out the gel carefully.
2. Submerge the gel in a tray containing freshly prepared 0.2 M HCl. Agitate gently for 10 min at room temperature to depurinate the DNA samples from the gel. The bromophenol blue dye in the gel should gradually turn grey during this process.
3. Rinse the gel three times with distilled water.
4. Submerge the gel in a tray of denaturing buffer, and agitate gently for 1 h at room temperature. The loading dye in the gel should turn back to blue during this process.
5. Rinse the gel three times with distilled water.
6. Submerge the gel in a tray of neutralization buffer, and agitate gently for at least 1 h at room temperature.
7. Soak the Hybond-XL membrane (12×10cm) in a tray with de-ionized water for 5 min at room temperature. Replace the water with 20× SSC and continue to agitate for 15 min.
8. To transfer DNA from the gel to the Hybond-XL membrane, assemble the transfer apparatus as following: layer two sheets of Whatman 3 MM filter paper in the gel transfer tray and pour 20× SSC transfer buffer on top to completely wet the Whatman papers. Smooth out the bubbles in between the Whatman papers and the tray gently by a plastic roller, and get rid of the excess amount of the transfer buffer. Place the gel on top of the Whatman papers with the side containing the loading wells facing down; make sure there is no air bubble between the Whatman papers and the gel. Place the presoaked Hybond-XL membrane on top of the gel and use a plastic roller to get rid of the air bubbles between the gel and the membrane. Layer two sheets of 20× SSC pre-wet Whatman papers (12×10cm) on top of the membrane, use plastic roller to remove air bubbles in between. Place a stack of dry paper towels (about 4 inches) pre-cut to the gel size on top of the Whatman papers. Seal the transfer tray with plastic wrap to prevent buffer evaporation during transfer. Put some weight (approx. 500g. such as metal plate, casserole dish etc.) on top of the transfer apparatus (Fig.1). Let the assembled transfer apparatus sit still on a flat surface and transfer for 24-48 h.

Note: When assemble the Southern blot transfer apparatus, avoid possible short-circuits of capillary liquid flow. The DNA will not be efficiently transferred if the blot short circuits. Use parafilm strips to seal the edges of the gel.

9. After gel transfer, disassemble the blot apparatus and flip over the membrane with the gel still attached, and mark the position of each loading well on the membrane by a pencil. Wrap up the gel and paper towels by plastic wrap and discard.

Note: After transfer, the gel becomes thinner. Use the pencil tip to penetrate the loading wells and mark their positions on the membrane. This step is for labeling the DNA binding site of the membrane and tracking the running lanes with orientation.

10. Crosslink the Hybond-XL membrane in a UV crosslinker chamber with UV energy dosage at 120 mJ/cm2. The membrane can be directly subject to hybridization, or sandwiched between two sheets of Whatman filters and stored at -20°C.

 

Probe Preparation for HBV DNA Detection

1. Prepare rNTP (supplied with Riboprobe® system) solution by mixing 50 μL of each 10 mM rATP, 10 mM rGTP, 10 mM rCTP, and 0.2 mM UTP.
2. Add 4 μL of 5 × reaction buffer, 4 μL of rNTP solution prepared from the previous step, 2 μL of 100 mM DTT, 0.25 μg of SalI-linearized pGEM-HBV DNA template, 1 μL of RNasin and 1 μL of SP6 polymerase into a nuclease free tube. Adjust the volume to 14 μL with nuclease-free water before adding 6 μL of [α-32P] UTP into the mixture. Incubate the reaction at 37ºC for 1 h.

Note: Linearization of the plasmid pGEM-HBV by SalI is an optimal step to produce “run-off” transcripts derived from the HBV sequence only. Other restriction enzyme, which singly cuts the vector sequences downstream of SP6-HBV transcription cassette but does not leave a 3’ overhang, can also be used.

Note: All radioactive materials and procedures should be handled strictly by following the institutional laboratory safety regulation rules.

3. To remove the DNA template, add 1 μL of RNase-free RQ1 DNase to the reaction and incubate at 37°C for 15 min.
4. Add 15 μL of 5M NH4OAc to stop the reaction. To precipitate the RNA probe, add 113 μL of nuclease-free water, 4 μL of yeast RNA, and 150 μL of isopropanol, gently mix and incubate at room temperature for 10 min.
5. Centrifuge the mixture at 12,000× g for 30 min at 4°C, and discard the supernatant carefully without disturbing the pellet. Dissolve the probes in 400 μL of de-ionized formamide, followed by measurement of the counts per minute (CPM) of acid insoluble 32P with scintillation counter (PerkinElmer). Store the probes at -20°C.

Note: 32P-radiolabeled HBV DNA probes prepared by random priming or end labeling can also be used in the hybridization.

 

Hybridization

1. Place the crosslinked membrane in a hybridization tube with the DNA-binding side facing the center of the tube. Add 5 mL of EKONO™ hybridization buffer, and pre-hybridize the membrane by rotating the hybridization tube at 65°C for 1 h in a hybridization oven.

Note: If using other commercial hybridization buffers, follow the pre-hybridization and hybridization conditions recommended by the manufacturers.

2. Replace the pre-hybridization buffer with 5 mL of fresh EKONO™ hybridization buffer, and add HBV riboprobes with 1×10⁷ Rotate the hybridization tube at 65°C overnight.
3. Discard the hybridization solution on the following day, and wash the hybridization membrane with approximately half a tube of wash buffer. Rotate at 65°C for 30 min.
4. Discard the wash buffer, and replace with half a tube of fresh wash buffer. Continue to wash at 65°C for 1 h.
5. After the second wash, take out the membrane and dry it with paper towels, making sure the membrane is not cracked or wrinkled during this process. Seal the membrane with plastic wrap. Place the membrane in a phosphorimager cassette with the DNA-binding side facing up. Layer the phosphor imaging screen on top facing the DNA-binding side of the membrane. Close the cassette tightly and expose overnight in the dark.
6. Scan the phosphor imaging screen with phosphorimager system. Store the membrane properly and erase the signal in the phosphor imaging screen with intense light for re-use.
7. A typical phosphor image of HBV DNA Southern blot is shown in Fig. 2 (also see in references (7, 9)). Quantify the signal intensity of the cccDNA bands with software provided by the phosphorimager system.

 

Figure 1:  Assembly of HBV DNA Southern blot transfer apparatus. See text for details.

Figure 2:  Detection of HBV DNA by a Southern blot assay. A typical pattern of HBV cytoplasmic core DNA (lane 2) and Hirt DNA (lane 3) upon Southern blot hybridization is shown. RC: relaxed circular DNA; DP-RC: deproteinized RC DNA; CCC: covalently closed circular DNA; SS: single-stranded (-) DNA. M: HBV genomic-length DNA marker. The approximate size of each HBV DNA species is labeled on the left. To further validate the authenticity of HBV cccDNA, the Hirt DNA sample in lane 3 has been heated to 85°C for 5 min before gel loading, a condition that denatures DP-rcDNA into SS DNA, while the cccDNA stays undenatured and its electrophoretic mobility remains unchanged (lane 4). The DNA sample from lane 4 is further digested with EcoRI, in which condition the cccDNA is linearized to a genome-length double-stranded DNA (lane 5). Reproduced from (9) with permission from Elsevier.