Here are a few protocols that may be of some help .....
Constructing a baculovirus Transfection 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 28C. Meanwhile prepare DNA: 2-4ug Transfer vector +5uL Baculogold DNA. Mix and incubate 5min at RT. 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. Aspirate medium form the attached insect cells, replace with 0.75 mL fresh medium. Gently pipet DNA mix onto cells, drop-by-drop, and gently spread to cover bottom of flask. Incubate 4h at 28C. Aspirate transfection mix. Rinse with 5mL fresh medium, aspirate again. Add 6mL fresh to each flask and return to incubator. Plaque assays 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. Spin 5min at 800rpm to pellet cells, debris (no need to transfer to new tube). 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). 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) Incubate 1h at 28C, swirling gently every 10min. Meanwhile, pre-warm 40mL of medium at 37C in sterile 50mL Falcon tube. 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). Add 40 mL pre-warmed medium to bottle and swirl vigorously to mix. Allow agarose/medium mixture to cool~5min (to ~37C). Aspirate virus from cells. Slowly pipet 4mL of agarose onto each plate, swirling to spread evenly. Quickly transfer to 28C incubator to set. 5days later, add trypan blue: Add agarose/medium mixture 1mL of sterile 1% trypan blue (or 5mL of premixed 0.4% trypan blue solution from Sigma) and swirl vigorously. Add 2mL of trypan blue/agarose/medium to each plate and quickly replace in incubator. 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! Prepare and label sterile Nunc cryotubes containing 1mL of medium each-as many tubes as you have plaques. 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). Transfer plaque to tubes. Pipetting medium up and down to eject all the agarose. Agitate tubes briefly (can vortex) and incubate >24h at 4C to allow virus to diffuse out the agarose plug. Amplification of Pass 1 viral stocks 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. Aspirate medium and replace with 0.9 mL of virus suspension (i.e. the plaque eluates). Incubate 1h at 28C, gently rocking every 10min or so to spread virus. Add 4mL fresh medium and incubate 3 days at 28C. 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). Testing expression 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 and positive control). Allow cells to attach ~30min and aspirate medium. Add 0.25 mL Pass 1 virus (or controls; medium [-] or virus [+]) to wells. Incubate 1h at 28C, gently rocking every 10min or so to spread virus. Add 1.5mL fresh medium and incubate 2 days at 28C. 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. Spin 5 min at 800rpm, remove and discard supernatant. 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-chilled Eppendorfs. Vortex (hard); add 5M NaCl to bring to total concentration to 150mM 3mL/100mL and repeat vortexing. Spin 20min at 14 000rpm at 4C (cold room microfuge). Remove supernatant and transfer to new pre-chilled Eppnedorf. Check protein concentration by Bradford (aka Bio-Rad) assay and load 50uL protein on SDS-PAGE gel. Stain with Coomassie blue and look for novel protein band by comparison with [+] control virus. Comparison with [-] no virus control is less helpful, as there are several bands that are induced by infection, regardless of foreign protein expression. Amplifying a Pass 2 virus stock Identify you r best-expressing Pass 1 virus for each virus you are making. Transfer 4x107 cells (split the day before) to sterile 50mL falcon tubes. You will need one tube for each virus you are amplifying. Spin 5min at 1500rpm, remove and discard supernatant. Resuspend cell in 3.5mL fresh medium. Add 0.5mL of Pass 1 virus; swirl to mix. Incubate 1h at 28C, gently swirling every 10min or so to mix cells and virus. Add 36mL fresh medium to each tube, mix well (but gently) and divide 20mL each into two T150 T.C. flasks. Incubate 3 days at 28C. 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.
Bac-To-Bac Baculovirus expression systems Methods: Transposition Prepare Luria Agar plates containing: 250mL -50mg/mL kanamycin 250mL from 50mg/mL -7mg/mL gentamicin 175mL from 10mg/mL 10mg/mL tetracycline 250mL from 10mg/mL 200mg/mL Bluo-gal 2.5mL from 20mg/mL 40mg/mL IPTG 50mL from 200mg/mL Thaw the DH10Bac competent cells on ice. Dispense 50mL of cells into 15mL polypropylene tubes. Add app. 1ng recombinant donor plasmid (in 5uL) and gently mix the DNA into the cells by tapping the side of tube. Incubate the mixture on ice for 30min. Heat shock the mixture by transferring to 42C water bath for 45sec. Chill the mixture on ice for 2min. Add 900uL S.O.C. medium to the mixture. Place the mixture in a shaking incubator at 37C with medium agitation for 4h. Serially dilute cells using S.O.C. medium to 10-1,10-2,10-3, 10-4 (i.e. 100mL of transposition mix: 900mL of S.O.C. medium=10-1 dilution, use this to further dilute 10-fold to give 10-2 dilution, and similarly for 10-3 and 10 -4 dilution). Place 100mL of each dilution on the plates and spread evenly over the surface. Incubate for at least 24h at 37C (Colonies are very small and blue colonies may not be discernible prior to 24h). Isolation of Recombinant Bacmid DNA White colonies contain the recombinant bacmid, and therefore, are selected of isolation of recombinant bacmid DNA. Before isolating DNA, candidate colonies are streaked to ensure they are truly white. Select white colonies from a plate with app. 100 to 200 colonies. Note: This number facilitates differentiation between blue and white colonies. Pick ~10 white candidates and streak to fresh plates to verify the phenotype. Incubate overnight at 37C. From a single colony confirmed as having white phenotype on plates containing Bluo-gal and IPTG, set up a liquid culture for isolation of recombinant bacmid DNA. The following protocol was specifically developed for isolating large plasmids (>100kb) and was adapted for isolating bacmid DNA. Using a sterile needle, inoculate a single, isolated bacterial colony into 2mL LB medium supplemented with 50ug/mL kanamycin, 7ug/mL gentamicin, and 10ug/mL tetracycline. A 15-mL snap-cap polypropylene tube is suitable. Grow at 37C to stationary phase (up to 24h) shaking at 250 to 300rpm. Transfer 1.5mL of culture to a 1.5-mL micro centrifuge tube and centrifuge at 14 000xg for 1min. Remove the supernatant by vacuum aspiration and resuspend (by gently vortexing or Pipetting up and down, if necessary) each pellet in 0.3 mL of Solution I (15mM Tris-HCl pH 8.0, 10mM EDTA, 100mg/mL RNase A). Add 0.3 mL of Solution II (0.2N NaOH, 1% SDS) and gently mix. Incubate at room temperature for 5min. Note: The appearance of the suspension should change from very turbid to almost translucent. Slowly add 0.3 mL of 3M Potassium acetate pH 5.5, mixing gently during addition. A thick white precipitate of protein and E.coli genomic DNA will form. Place the sample on ice for 5 to 10 min. Centrifuge for 10min at 14 000 x g. During the centrifugation, label another micro centrifuge tube and add 0.8 mL absolute isopropanol to it. Gently transfer the supernatant to the tube containing isopropanol. Avoid any white precipitate material. Mix by gently inverting tube a few times and place on ice for 5 to 10 min. At this stage, the sample can be stored at -20C overnight. Centrifuge the sample for 15min at 14 000 x g at room temperature. Remove the supernatant and add 0.5mL of 70% ethanol to each tube. Invert the tube several times to wash the pellet. Centrifuge for 5min at 14 000 x g at room temperature. (Optional: repeat step 8) Remove as much of the supernatant as possible. Note: The pellet may become dislodged from the bottom of the tube, so it is better to care fully aspirate the supernatant than to pour it. Air-dry the pellet briefly, 5 to 10 min, at room temperature and dissolve the DNA in 40uL TE. Allow the solution to sit in the tube with occasional gentle tapping of the bottom of the tube. The DNA is generally ready for use within 10min, as long as the pellets are not over dried. Store the NA at -20C. However, avoid repeated freeze/thaw cycles to avoid a drastic reduction in transfection efficiency. Preparations of bacmid DNA may be analyzed by agarose gel electrophoresis to confirm the presence of high molecular weight DNA. Transfection of Sf9 Cells with Recombinant Bacmid DNA Seed 9x105 cells per 35-mm well (of a 6-well plate) in a 2mL of Sf-900 II SFM containing penicillin/streptomycin at 0.5X final concentration (50units/mL penicillin, 50mg/mL streptomycin). Use only from a 3- to 4- day-old suspension culture in mid-log phase with viability of >97%. Allow cells to attach at 27C for at least 1h. Prepare the following solutions in 12 x 75-mm sterile tubes: Solution A: For each transfection, dilute ~5mL of mini-prep bacmid DNA into 100mL Sf-900 II SFM without antibiotics. Solution B: For each transfection, dilute~6mL CellFectin Reagent into 100mL Sf-900 II SFM without antibiotics. Mix thoroughly by inverting the tube 5-10 times before removing a sample for transfection to ensure that a homogeneous sample is taken. Combine the two solutions, mix gently and incubate for 15 to 45 min at room temperature. Wash cells once with 2mL of Sf-900 II SFM without antibiotics. For each transfection, add 0.8 mL of Sf-900 II SFM to each tube containing the lipid-DNA complexes. Mix gently. Aspirate wash media from cells and overlay the diluted lipid-DNA complexes onto cells. Incubate cells for 5h in a 27C incubator. Remove the transfection mixtures and add 2mL of Sf-900 II SFM containing antibiotics. Incubate cells in a 27C incubator for 72h. Harvest virus from cell culture medium at 72h post-transfection. Harvest/Storage of Recombinant Baculovirus When harvesting virus from the transfection, transfer the supernatant (2mL) to a sterile, capped tube. Clarify by centrifugation for 5min at 500 x g and transfer the virus containing to a fresh tube. From the initial transfection, viral titers of 2x107 to 4x107 pfu/mL can be expected. Store the virus at 4C, protected from light. For long tem storage of virus, the addition of fetal bovine serum (FBS) to a final concentration of at least 2%FBS is recommended. Storage of an aliquot of the viral stock at -700C is also recommended. For determining the viral titer, a plaque assay can be performed. Fro amplifying viral stocks, infect a suspension or monolayer culture at a Multiplicity of Infection (MOI) of 0.01 to 0.1. Use the following formula: Innoculum required (mL): desired MOI (pfu/mL) x (total number of cells) Titer of viral innoculum (pfu/mL) For example, infect a 50-mL culture at 2 x 106 cells/mL with 0.5 mL of a viral stock that is 2x107 pfu/mL, for another MOI of 0.1. Harvest virus at 48h post-infection. This will result app. 100-fold amplification of the virus. Preparation of Stock Solutions Antibiotics can be ordered in either dry powdered from or as a stabilized, sterile, premixed solution. These solutions should be stored according to the manufacturer's recommendations. Stock solutions of antibiotics dissolved in water should be sterilized by titration trough a 0.22-micron filter. Antibiotics dissolved in ethanol need not be sterilized. Store stock solutions in light-tight containers. Magnesium ions are antagonists of tetracycline. Use media without magnesium salts for selection of bacteria resistant to tetracycline. Antibiotic Stock solution concentration Storage Ampicillin 50mg/mL in water -20C Kanamycin 10mg/mL in water -20C Tetracycline 10mg/mL in ethanol -20C Gentamicin 7mg/mL in water -20C Bluo-gal solutions are made by dissolving the dry powder in dimethylformamide or dimethyl sulfoxide (DMSO) to make 20 mg/mL stock solution. Care must be taken when using dimethylformamide, dispense solution in a vented chemical hood only. Use glass or polypropylene tube. The tube containing the solution should be wrapped in aluminum foil to prevent damage by light and stored at -200C. It is not necessary to sterilize the solution by filtration. A 200mg/mL stock of IPTG is made by mixing 2g of IPTG with 8mL of water until dissolved. Adjust the volume of the solution to 10mL water and sterilize by filtration through a 0.22-micron filter. Dispense the solution to 10mL with water and sterilize by filtration trough a 0.22-micron filter. Dispense the solution into several 1-mL aliquots and store at -20C.
Titering Virus-Protocol Plate 2x106 cells in Lux plates in total of 3mL (e.g. 1mL Sf9 at 2x106 and 2mL medium). Allow to attach, then aspirate supernatant. For each virus make a 10-5, 10-6, 10-7 dilution in Grace's complete e.g. make 1:100 dilution (5mL in 5 mL), then dilute 1:100 (50mL in 5mL) 10-5 dilute 1:10 (500mL in 5mL) 10-6 dilute 1:10 (500mL in 5mL) 10-7 Pipet 1mL of each dilution onto a separate dish of cells. Incubate for 1h at 28C, agitating by rocking every 10 in, then aspirate virus. Meanwhile, pre-warm 40mL medium to 37C. Melt 5% LMP-agarose, mix well. Allow to cool to app, 28-37C, then carefully pipet 4mL per dish onto cells. Incubate at 28C for 5-6 days. On day 5 or 6 make 1% LPM-agarose/grace's as before, but add to it 1mL 1% Trypman blue, mix and add 2mL to each plate. In 1 or 2 days, count plaques.
Infecting Sf9 cells on plates Plate 25x106 cells (split the day before) per 150mm T.C. dish in a total of 15mL(e.g. 12.5 mL cells at 2x106 cells/mL plus 2.5mLmedium). Allow cells to attach 30-60min. Aspirate medium. Add virus to attached cells. Want m.o.i. of 5, or 1.25x108 pfu for 25x106 cells. For typical Pass 2 or 3 stocks with a titer of 108pfu/mL, therefore use 1.25mL virus per dish. If, in a pinch, you need to use an untitred stock, double the volume, just to be safe. Rock plate gently to spread virus over cells. Incubate 1h at 28C, rocking dishes every 10 min or so to spread the virus. Add 10mL fresh medium to each plate and incubate 48h at 28C. 2 days after infection, harvest cells by gently scraping with rubber policeman, to resuspend in medium (do not attempt to wash cells on plate). Keep cells submerged as you scrape to avoid lysis. Transfer suspended cells to 15mL conical tubes, which you may top off with HBS. Spin 5-10 min at 1000rpm, aspirate supernatant. Resuspend cell pellet in 1mL per dish lysis buffer (see protocol for making Pass 1 lysate), pipet up and down to resuspend, and transfer to pre-chilled Eppendorf tubes. Vortex (hard) 15 sec, add 5M NaCl to bring total concentration up to 150mM, vortex 15 sec again. Spin 20 min at 14 000rpm in cold room microfuge. Collect supernatants and pool extracts form identical infections. This is your crude lysate. Check protein concentration of 2mL and 5mL. Desalting lysate (optional). Th8is step is to remove small molecules, nucleotides, etc. and seems to help reduce the background of CDK activation seen in crude insect cell lysates. Equilibrate PD10 desalting column (Pharmacia) in 25mM Hepes, pH 7.4; 150mM NaCl; 1mM EDTA; 1mM DTT; 10% glycerol. Note: This is a standard FPLC column buffer, which you can make by mixing appropriate Buffers A (0 salt) and B (1M NaCl) in an 85:15 volume ratio. Follow Pharmacia's directions for equilibrating and loading PD 10 columns exactly. You will end up with 3.5mL of desalted lysate. Re-check protein concentration.
Baculovirus Amplification (Pass 2 or Small Scale Pass 3) Remove 4x107 cells to a 50mL Falcon tube (~20mL at 2x106). Pellet at 1500rpm 5min. Aspirate supernatant. Resuspend in 0.5mL virus (at1x108) with 3.5mL fresh medium. MOI=0.5-1.0. Incubate 60min at 28C with occasional swirling. Add 36mL medium, mix and plate 20mL each on T-150 flasks. Incubate 3-4 days at 28C. Harvest supernatant and spin down cells and debris. Baculovirus Amplification (Pass 3 or Large Scale) Divide 150mL of cells at 2x106 (3x108 total) into three 50mL Falcon tubes. Pellet cells at 1000rpm for 8min. Aspirate supernatant. Resuspend in Pass 2 virus at MOI of 0.5 ~1.5mL at 1x108 for all the three tubes. Add 9x virus volume of fresh medium (~13.5mL). Incubate at 28C for 1h with occasional swirling. Transfer mix to fresh flask with 300mL of fresh medium. Incubate 3-4days at 28C. Harvest to six 50mL Falcon tubes Spin 10min at 1000rpm.Store at 40C. Note: If the supernatant is cloudy check by microscopy for contamination.