Constructing a baculovirus, 1997

  1. Transfection

    1. Need Sf9 insect cells, growing logarithmically at ~2x106 cells/ml; plate 2x106 cells in T25 T.C. flask along with 4ml medium (complete Grace's +10% FBS, 1x Pen /Strep, 0.1% Pluronic). Allow cells to attach for ~30min at 280C.

    2. Meanwhile prepare DNA:

2-4mg Transfer vector +5ml Baculogold DNA. Mix and incubate 5min at R.T. Add 0.7 ml Transfection buffer:

25mM Hepes-NaOH, pH 7.1

140mM NaCl

125mM CaCl2

(Make 100ml, re-check pH and adjust if necessary, then sterile-filter, using disposable filter apparatus in T.C. hood)

Mix gently by pipetting up and down.


    1. Aspirate medium form the attached insect cells, replace with 0.75 ml fresh medium

    2. Gently pipet DNA mix onto cells, drop-by-drop, and gently spread to cover bottom of flask

    3. Incubate 4h at 280C

    4. Aspirate transfection mix

    5. Rinse with 5ml fresh medium, aspirate again

    6. Add 6ml fresh to each flask and return to incubator


2. Plaque assays

  1. 5 days (can be 6 or 7) post-transfection, harvest virus in transfection supernatant by carefully removing with sterile pipet and transferring to sterile 15ml conical tube.

  2. Spin 5min at 800rpm to pellet cells, debris (no need to transfer to new tube).

  3. Plate 2x106 cells (split the day before) together with 3 ml medium in 60mm Lux plates, allow 30min to attach and aspirate medium (will need 4 plates per virus being constructed).

  4. Add to attached cells 1ml of virus suspension, diluted as follows:

Undiluted transfection supernatant (10 0)

Transfection sup. Diluted 1/10 (10 -1)

" " " 1/1000 (10-3)

" " " 1/100 000 (10-5)

  1. Incubate 1h at 280C, swirling gently every 10min

  2. Meanwhile, pre-warm 40ml of medium at 370C in sterile 50ml Falcon tube

  3. With ~5-10min to go in 1h incubation. Microwave 1 bottle of autoclaved LMP agarose (0.5mg/10ml water) to completely melt it (~1min on high)

  4. Add 40 ml pre-warmed medium to bottle and swirl vigorously to mix.

  5. Allow agarose/medium mixture to cool~5min (to ~370C).

  6. Aspirate virus from cells.

  7. Slowly pipet 4ml of agarose onto each plate, swirling to spread evenly.

  8. Quickly transfer to 280C incubator to set.

5days later, add trypan blue

  1. Follow steps 2.f-I above

  2. Add agarose/medium mixture 1ml of sterile 1% trypan blue (or 5ml of premixed 0.4% trypan blue solution from Sigma) and swirl vigorously.

  3. Add 2ml of trypan blue/agarose/medium to each plate and quickly replace in incubator.

3.Picking plaques

1-2 days after adding trypan blue, plaques should be clearly visible. Look under microscope (low power) at what you think is a plaque by eye-you should see clearly cells that are blue (dead) surrounded by cells that are clear (alive). For each virus you are trying to isolate, find 4-6 plaques that are well separated from neighboring plaques (you may have to use more than one dilution), circle them with a marking pen, check each one under the scope to ensure that it's real plaque. Now the fun begins.

  1. Prepare and label sterile Nunc cryotubes containing 1ml of medium each-as many tubes as you have plaques.

  2. Place Lux plate on light box, tubes in T.C. hood. Using sterile yellow tip on P200, carefully excise plaque from agarose by plunging tip down to the plastic with plunger depressed, then simultaneously, slowly withdrawing tip and drawing back the plunger. The agarose plug containing the plaque should squirt into the yellow tip (check the "cored" agarose under microscope if you like, to make sure you didn't miss)

  3. Transfer plaque to tubes. Pipetting medium up and down to eject all the agarose.

  4. Agitate tubes briefly (can vortex) and incubate >24h at 40C to allow virus to diffuse out the agarose plug.

4. Amplification of Pass 1 viral stocks

  1. Plate 2x106 cells (split the day before) in T25 flasks with 4ml of medium allow to attach ~30min-need one flask per plaque you picked.

  2. Aspirate medium and replace with 0.9 ml of virus suspension (i.e. the plaque eluates)

  3. Incubate 1h at 280C, gently rocking every 10min or so to spread virus

  4. Add 4ml fresh medium and incubate 3 days at 280C.

  5. Harvest virus by carefully removing with a sterile pipet, transfer to 15ml conical tube and spin 5min at 800rpm to pellet cells and debris (no need to transfer to new tube)

5. Testing expression

  1. Plate 1x106 cells (split the day before), together with 2ml of medium, in 6 well dishes (1 well per Pass 1 stock you are testing, plus negative [no virus] and positive [high expressing virus such as cyclin H] control). Allow cells to attach ~30min and aspirate medium.

  2. Add 0.25 ml Pass 1 virus (or controls; medium [-] or virus [+]) to wells.

  3. Incubate 1h at 280C, gently rocking every 10min or so to spread virus.

  4. Add1.5ml fresh medium and incubate 2 days at 280C.

  5. Prepare lysates (need no longer be sterile). Scrape cell gently (to avoid premature lysis) off the bottom of dish, into medium and transfer to 15ml conical tubes. Top tubes off to ~10ml each with HBS 910mM Hepes, pH 7.4; 150mM NaCl).

  6. Spin 5 min at 800rpm, remove and discard supernatant.

  7. Resuspend cell pellet in 100ml of lysis buffer:

For 100ml

10mM Hepes-NaOH, pH 7.4 1ml 1M

10mM NaCl 200ml 5M

5mM EDTA 1ml 0.5M

1mM DTT*

0.5 mM PMSF*

2mg/ml aprotinin*

1mg/ml leupetin*

*add fresh just prior to use

Pipet up and down to resuspend and transfer to pre-chilled1.4ml Eppendorrfs.

  1. Vortex (hard); add5M NaCl to bring to total concentration to 150mM 3ml/100ml) and repeat vortexing.

  2. Spin 20min at 14 000rpm at 40C (cold room microfuge)

  3. Remove supernatant and transfer to new pre-chilled Eppnedorf. Check protein concentration by Bradford (aka Bio-Rad) assay and load 50ml protein on SDS-PAGE gel. Stain with Coomassie blue and look for novel protein band by comparison with [+] control virus (may need Western if low expression and you have an antibody or tagged protein). Comparison with [-] no virus control is less helpful, as there are several bands that are induced by infection, regardless of foreign protein expression.

6. Amplifying a Pass 2 virus stock

  1. Identify you r best-expressing Pass 1 virus for each virus you are making

  2. Transfer 4x107 cells (split the day before) to sterile 50ml falcon tubes. You will need one tube for each virus you are amplifying.

  3. Spin 5min at 1500rpm, remove and discard supernatant.

  4. Resuspend cell in 3.5ml fresh medium.

  5. Add 0.5ml of Pass 1 virus; swirl to mix

  6. Incubate 1h at 28 0C, gently swirling every 10min or so to mix cells and virus.

  7. Add 36ml fresh medium to each tube, mix well (but gently) and divide 20ml each into two T150 T.C. flasks

  8. Incubate 3 days at 28 0C.

  9. Spin out debris, 10min at 1500rpm (no need to transfer to fresh tubes); label tubes "A" and "B" and store at 40C. These are your Pass 2 stocks, which are adequate for most small- to medium-scale infections. For large-scale purification, you will need to make Pass 3 stocks. All working stocks (Pass 2 or higher) should be tittered. If the titer is less than 108 plaque-forming units (pfu) per ml you may have problems and may need to re-amplify from a high titer stock as earlier passage.