Insect Expression Support—Troubleshooting

Please review the possible reasons and recommendations below:

• Incomplete digestion of pFastBac™ plasmid or insert DNA: Use additional restriction enzyme for digestion or purify the insert DNA.
• Incomplete or excessive phosphatase treatment of pFastBac™ plasmid: Optimize dephosphorylation conditions according to the manufacturer’s recommendations for the phosphatase you are using.
• Poor recovery of pFastBac™ plasmid or insert DNA from agarose gel: Use our PureLink® Quick Gel Extraction System to purify high-quality plasmid DNA from your agarose gel.
• Incomplete ligation reactions: Follow ligation conditions and optimize the reaction by varying vector:insert molar ratios (1:3, 1:1, 3:1).
• Insert contains unstable DNA sequences such as LTR sequences and inverted repeats: Grow transformed cells at a lower temperature (30°C) or try a different competent cell strain such as MAX Efficiency Stbl2™ Competent Cells.

Please review the possible causes and recommendations below:

• Low transformation efficiency of competent E. coli: If stored incorrectly, prepare or obtain new competent cells. We recommend using our One Shot® TOP10 or One Shot® MAX Efficiency® DH10B™-T1R chemically competent E. coli cells.
• Impurities in DNA: Purify the insert DNA. Ensure that any excess phenol, proteins, detergents, and ethanol from the DNA solution are removed.
• Too much DNA was transformed: For chemically competent cells, add 1–10 ng of DNA in a volume of 5 µL or less per 100 µL of cells. For electrocompetent cells, add 10–50 ng of DNA in a volume of 1 µL or less per 20 µL of cells.
• Incomplete ligation reaction: Optimize your ligation reaction and include a ligation control (i.e., digested pFastBac™ vector + ligase; no insert).
• Ligation reaction mix inhibits transformation of competent cells: Reduce the amount of ligation reaction transformed. Dilute ligation reaction 5x with TE buffer prior to transformation.
• Problem with antibiotic: Confirm that you are using the correct antibiotic and concentration, as well as the age of the antibiotic.
• Competent cells stored improperly or handled improperly: Store competent cells at –80°C; thaw cells on ice and use immediately; do not vortex.

Although you will be picking white (recombinant) colonies, you should expect to see some blue (contain non-recombinant bacmid) colonies. Here are some possible causes for seeing no blue colonies and recommendations for the same:

• Insufficient time for color development: Wait at least 48 hours before identifying colony phenotypes.
• Use Bluo-gal instead of X-Gal in agar plates: Bluo-gal helps to increase contrast between blue and white colonies.
• Insufficient growth after transposition: Grow transformed cells in SOC medium for a minimum of 4 hours before plating.
• Bluo-gal and IPTG omitted from plates: Prepare fresh selective plates containing 50 µg/mL kanamycin, 7 µg/mL gentamicin, 10 µg/mL tetracycline, 100 µg/mL Bluo-gal, and 40 µg/mL IPTG.
• There are too many colonies on the plate: Serially dilute the transformation mix to obtain well-spaced colonies (10-2 to 10-4 is suggested).
• Plates are too old or stored in light: Do not use plates that are more than 4 weeks old; store plates protected from light.
• Incubation period too short or temperature is too low: Wait at least 48 hours before picking colonies. Incubate plates at 37°C.

Please review the following possibilities and recommendations:

• pFastBac™ DNA used for transformation was of poor quality: Use purified plasmid DNA for transformation and check the quality of your plasmid DNA.
• Gentamicin omitted from plates:  Prepare fresh selective plates containing 50 µg/mL kanamycin, 7 µg/mL gentamicin, 10 µg/mL tetracycline, 100 µg/mL Bluo-gal, and 40 µg/mL IPTG.

Please review the following reasons and our recommendations:

• Use LB medium for recovery/expression period: Use SOC medium for the 4 hr growth time.
• Recovery/expression time too short: Increase the recovery time to >4 hr at 37°C or 6 hr at 30°C.
• IPTG concentration is not optimal: We suggest using 20–40 µg/mL IPTG.

We recommend analyzing your recombinant bacmid DNA by PCR analysis. Use the pUC/M13 Foward and Reverse primers that hybridze to sites flanking the mini-attTn7 site within the lacZalpha-complementation region to facilitate PCR analysis. Please see page 32 of the manual for further instructions.

Poor color differentiation for your colonies could be caused by the following:

• Agar is not at the correct pH: Adjust pH of LB agar to 7.0.
• Intensity of the blue color is too weak; ensure that you are using Bluo-gal, not X-Gal. You can also try increasing the concentration of Bluo-gal to 300 µg/mL.
• Too many or too few colonies on the plate: Adjust the serial dilutions of cells to obtain an optimal number of colonies. 
• Incubation period too short or temperature too low: Do not pick colonies until 48 hours after plating; incubate plates at 37°C.
• IPTG concentration is not optimal: A range of 20–60 µg/mL IPTG generally gives optimal color development.

This could be caused by the following:

• Wrong antibiotic or old media: use fresh media.
• Colonies are too old or too small: Use large white colonies from freshly streaked plates.
• Unstable insert caused by special feature of the gene of interest; for example, direct repeats: Incubate the culture at 30°C for 24 hours instead of 37°C overnight.

Please see the possible reasons and suggestions below:

• DNA stored improperly: Ensure that purified bacmid DNA is stored at –20°C in aliquots to avoid repeated free/thaws.
• High molecular weight bacmid DNA handled improperly: When isolating bacmid DNA, do not vortex the DNA solution; additionally, do not resuspend DNA pellets mechanically; allow solution to sit in the tube with occasional tapping.

Please see the possible causes and suggestions we have to alleviate this problem:

• Use pure proofreading polymerase for analysis: Use a Taq-based polymerase for the analysis.
• Insert is very long and causes difficulties in PCR: Instead of using both M13 forward and reverse primers, use one gene-specific primer paired with the M13 primer of your choice.
• Long GC-rich stretches in the gene of interest: Consider using DMSO (up to 8%) in the PCR reaction.

Most likely, a colony that was gray or dark in the center was picked. Try to analyze more white DH10Bac™ transformants. Typically, we recommend picking a white colony whose diameter is >2 mm. Restreak the white colonies on a fresh plate with 50 µg/mL kanamycin, 7 µg/mL gentamicin, 10 µg/mL tetracycline, 100 µg/mL Bluo-gal and 40 µg/mL IPTG. Incubate plates for 24 hours.

This may be due to contamination or cytotoxicity from the bacmid prep. Make sure to include a negative control that is the bacmid only without Cellfectin® II Reagent. Additionally, use the PureLink® HiPure plasmid prep kit, not the silica-based miniprep kit for bacmid prep.

There are several possibilities:

• Using media containing antibiotics during transfections.
• Plating cells at too low a density: We recommend at least 70% confluence.
• Using cells at too early a passage: We recommend growing cells for at least 5 passages before using them for transfection.
• Contamination because of no pen/strep after the transfection: After 5–8 hr incubation with the transfection mixture, remove the mixture and add antibiotics containing media/well. 

Please see the possible reasons and suggestions below:

• Mixture of Cellfectin® II Reagent and bacmid was not performed or was not incubated long enough: Mix the Cellfectin® II Reagent and bacmid well by tapping or gentle vortexing, and incubate the mixture for 15–45 min.
• Bacmid yield is lower than estimated: Set up an optimization with different amounts of bacmid.
• Bacmid is sheared during purification or freeze/thaw: Verify the integrity of bacmid on a gel.
• Incubation time is not long enough: Incubate mix for 8 hr at 27°C.
• Cells used are of high passages or have passed log-phase growth: For best results, use cells between 8–15 passages; plate cells when they are in log-phase growth.
• Cellfectin® II Reagent has been frozen: Purchase a new vial.
• Medium used contains serum: Use unsupplemented Grace’s medium in transfection.

Check the MOI. It may be low because the titer of the P1 virus is lower than what was estimated.

Please see the following reasons and suggestions to improve yield:

• Low transfection efficiency: We recommend using our Cellfectin® II Reagent for transfection; perform the transfection in Grace’s Medium, Unsupplemented and ensure no supplements, FBS, or antibiotics are present during the transfection. Harvest the viral supernatant when signs of infection are visible (typically >96 hr post-transfection).
• Cells are plated too sparsely; check recommended cell densities.
• Use too much or too little Cellfectin® II Transfection Reagent: Optimize amount used.
• Time of incubation with DNA:lipid complexes is too short or too long: Optimize incubation time (3–8 hr).
• Recombinant bacmid DNA is degraded: Check the quality of your recombinant DNA by agarose gel electrophoresis prior to transfection; prepare bacmid DNA using PureLink® HiPure Plasmid DNA Miniprep or Maxiprep Kit.
• Bacmid DNA is not pure (contains recombinant and empty bacmid): Screen other DH10Bac transformants and perform plaque purification to isolate recombinant baculovirus.

Low protein yield may occur due to the following reasons:

• Viral stock contains a mixture of recombinant and non-recombinant baculovirus: Perform plaque purification to isolate recombinant baculovirus.
• Baculovirus is not recombinant: Verify transposition by PCR analysis of bacmid DNA using pUC/M13 forward and reverse primers; re-transfect insect cells with new recombinant bacmid DNA.
• Use too low or too high viral titer: Vary the MOI.
• Time of cell harvest is not optimal: perform a time course of expression to determine the optimal time to obtain maximal protein expression.
• Cell growth conditions and medium are not optimal: Optimize cell culture conditions based on the size of your culture vessel and expression conditions; we recommend using Sf-900™ II SFM or Sf-900™ III SFM for optimal cell growth and protein expression.
• Cell line is not optimal; try other insect cell lines.
• Cells were harvested too late: Do a time-course experiment and harvest cells at different time points.

This can happen when:

• A binding partner or other parts of the protein are needed for folding: Identify the partner and perform coexpression of both proteins.
• Protein is naturally secreted, but is now expressed as intracellular: Express the protein as secreted.
• Protein is not extracted properly: For complete extraction, use sonication and DNase I. Keep samples cool at all times.
• Soluble tag may be needed: Consider using an N-terminal GST tag.

MOI and harvesting time need to be tuned after scale-up.

Please review the following possibilities and solutions:

• Viral stock was amplified using high MOI originally: Go back to the lower-passage viral stock and do a low-MOI amplification.
• Did not spin down and get rid of cells when harvesting viral supernatant: Go back to the lower-passage viral stock and do a low-MOI amplification; if this viral stock is P2, this stock can be used in amplification.
• For some genes, the virus can become very unstable: Free the aliquoted P2 viral stock and do one run of amplification after reviving.

Media used to culture insect cells usually have an acidic pH (6.0–6.5) or contain electron-donating groups that can prevent binding of the 6xHis-tagged protein to Ni-NTA. Amino acids such as glutamine, glycine, or histidine are present at significantly higher concentrations in media for growing insect cells than in media for growing mammalian cells, and compete with the 6xHis-tagged protein for binding sites on Ni-NTA matrices. Grace’s medium (Life Technologies), for example, contains approximately 10 mM glutamine, 10 mM glycine, and 15 mM histidine.

Dialysis of the medium against a buffer with the appropriate composition and pH (8.0) similar to the lysis buffer recommended for purification under native conditions usually restores optimal binding conditions. Note that depending on the medium used, a white precipitate (probably made up of insoluble salts) can occur, but normally the 6xHis-tagged protein remains in solution. This can be tested by either protein quantitation if using a protein-free medium or by monitoring the amount of 6xHis-tagged protein by western-blot analysis. After centrifugation, 6xHis-tagged protein can be directly purified from the cleared supernatant.

If you were using SF-900™ II SFM, that is incompatible with the ammonium acetate precipitation method. Our SF-900™ II Serum-Free Medium contains the block co-polymer non-ionic surfactant Pluronic F-68, which has been found to interfere with ammonium acetate precipitation. If you are putting the un-concentrated supernatant over the column, this should not cause a problem. If the supernatant has been concentrated, the Pluronic acid will need to be removed using a column. 

Warm the ganciclovir solution in a water bath at 37°C for 5-10 min, then vortex for a few minutes. The precipitate should go back into solution.

Please see our recommendations below:

• Check the LR reaction by PCR analysis prior to transfection into insect cells.
• We recommend using Grace’s Insect Cell Culture Medium, Unsupplemented during the transfection experiment instead of serum-free medium, as components in serum-free medium may interfere with transfection.
• Ensure that FBS, supplements, or antibiotics are not included during transfection, as the proteins in these materials can interfere with the Cellfectin® II Reagent.
• Use the LR recombination reaction using the pENTR/CAT plasmid as a positive control and Cellfectin® II Reagent only (mock transfection) as a negative control.
• Ensure that cells are in the log phase of growth with >95% viability, and the amount of cells are in accordance with the suggestions in the manual.
• Cells may not show signs of viral infection for up to a week depending on transfection efficiency; continue culturing and monitor cells daily for signs of infection.

To get a high-titer stock, reinfect cells with the P1 stock and generate a P2 high-titer stock. Follow the directions in the BaculoDirect™ manual on page 18 to generate your P2 stock.

Please check the construction of your entry clone, and ensure that the insert is in frame with the vector. Analyze the recombinant viral DNA by PCR to confirm the correct size and orientation of your insert after the LR reaction. Sequence your PCR product to verify the proper reading frame for expression of the epitope tag. 

Please see the following suggestions:

• The incorrect MOI was used; ensure that the amount of viral stock was calculated correctly, and that an MOI of 5–10 was used. You may need to test a range of MOIs depending on the kinetics of expression of your recombinant protein.
• The protein may be lost during cell lysis; if you are trying to detect an intracellular protein, analyze the supernatant to determine if the protein is being lost due to cell lysis.
• The protein is being degraded or unstable; add protease inhibitors to your cell lysates and/or check mRNA levels.
• The protein of interest is toxic to the cells; harvest the cells at earlier time points (e.g., 18–24 hr post-infection).

Please compare your cells-only plate to the infected plate. The uninfected cells should appear overgrown when compared to infected cells, as transfection inhibits growth. If this is seen, keep checking the infected cells daily for other signs of infection (nuclear swelling, detachment from the plate, viral budding, and lysis). The kinetics of infection may be slower than expected. If cell growth does not appear to be inhibited, you may consider the following factors:

• How was the DNA prepared? We recommend using a resin-based DNA purification system, such as our PureLink® HiPure Plasmid Prep Kits.
• Were cells in log phase? What was their confluency? We recommend that cells be in log phase, 95% viable, and plated at 50–70% confluency for successful transfection.
• What transfection reagent was used? We recommend using Cellfectin® II Reagent.

If there is contamination with wild-type virus in all of your samples, take a P1 viral stock and redo the plaque assay. Be sure to select well-spaced, occ– plaques. If you are having difficulty distinguishing a recombinant plaque from a non-recombinant plaque, try using one of the pBlueBac vectors. Recombinant plaques will be blue when chromogenic substrate is incorporated into the medium during the agarose overlay (see page 14 of the manual for more information).

An MOI of 5–10 is typically used. If too much virus is added, unfortunately the cells die too soon and the protein expression level goes down.

Too many cells were seeded; we recommend seeding 8 x 10⁵ cells per well for a 6-well plate.

Yes, this is indicative of an aspirating problem on the plaques. The agarose overlays were "floating" because the medium was not completely aspirated from the plates. The plates need to be completely dry before the agarose is placed over the cells, especially when plaques will be picked. To do this, we typically tip the plate slightly and keep going around the rim of the plate with the Pasteur pipette tip, being careful not to disturb the cell monolayer. If any medium pooling at the rims of the plates (they will be small pools) is seen, continue to aspirate. This “floating" agarose overlay problem may also result in wild-type contamination. The wild-type virus is able to migrate to other portions of the plates and contaminate recombinant plaques. Wild-type virus replicates much faster than recombinant virus, and can quickly overwhelm the recombinant virus.

The agarose overlay was too hot. After addition of the agarose overlay, cells should still be round and healthy.

There are a few things that can turn plates blue:

• Too much virus when plating. Try a higher dilution.
• Cells are being singed when plated with hot melted agarose. This lyses the cells and releases lacZ into the agarose, turning it blue. Double-check plating temperatures. If plates are too wet, the blue can diffuse.

On the day you intend to pick plaques, make a solution of Bluo-gal in DMSO at 20 mg/mL. Add 50 µL per plate and spread with a glass spreader under sterile conditions. Wait 30–60 min, and your plaques should turn blue.  

Normally, very small white dots show up about 5–7 days and 1 mm plaques show up around day 10. Plaques can vary in size from 1 mm to 4 mm.

Yes, cells are infected with wild-type virus individually and will develop polyhedra at different rates until all the cells in the flask are infected. The polyhedra in cells will form in approximately 3–4 days, differing in size and number until they reach their maximum capacity and burst the cell, releasing tiny particles of virus into the medium.

This is typically an indication of poor homologous recombination. Check the plasmid/linear DNA ratio you used. If there are some blue plaques, however, expand those viruses and check for their protein. In our experience, they are correct, even if they were in relatively low abundance.

In the case of a blue colony, the E. coli has the bacmid and the plasmid in it, allowing the cells to survive the selection process. However, because the transposition has not occurred, the LacZ gene is not disrupted. For bulls-eye colonies, this indicates that the transposition took place when the colony was growing. Re-streaking for an isolated clone from the white portion of the mixed colony should yield some colonies where transposition occurred.

The concentration of gentamicin might be too high. Try lowering the amount to 5 µg/mL and try adding more of the colony to the culture medium.

In instances where ExpiSf9 Cells are thawed and do not start to grow, or attain relatively low densities in culture, one common solution is to verify the culture volume and shake speed according to Table 1 in the ExpiSf Expression System Smart Start Guide.

If ExpiSf9 Cells significantly overgrow a density of 10 x 10E6 viable cells/mL during routine subculturing, we recommend passaging the cells down to a lower cell density of 0.5 x 10E6 - 1.0 x 10E6 viable cells/mL to reduce stress on the cells. Subculture cells a couple of times and monitor cell viability and growth kinetics to ensure that the cells have recovered and are healthy (i.e., ≥90% viability and reaching a viable cell density of ≥5 x 10E6 cells/mL 3-4 days post-passaging) before proceeding with experiments. If cells are allowed to overgrow above a density of 10 x 10E6 viable cells/mL on multiple occasions and growth patterns have been affected, we recommend establishing a new culture by thawing a new vial of cells.

The ExpiSf Enhancer is designed to work with the ExpiSf Expression System for maximal protein expression. If the enhancer is not added 18-24 hrs prior to infection with baculovirus, there will be a greater than 50% loss in protein expression.

We have observed a decrease in cell infectivity when the cell density is >7 x 10E6 cells/mL at the time of infection. If the cell density is >7 x 10E6 cells/mL 24 hours after addition of ExpiSf Enhancer, it is likely that the initial cell seeding density at Day 1 may have been above 5 x 10E6 cells/mL. We recommend verifying your cell counts using an alternative method, such as trypan blue exclusion using a hemocytometer, to ensure that cells are being accurately counted and seeded for your experiments.

An MOI of 5-10 is typically recommended for expression of most proteins with the ExpiSf Expression System. If too much virus is added, unfortunately the cells would die too soon and the protein expression level would go down.

The optimal complexation time is 5 mins. We have observed a gradual drop in protein yield if complexes sit for longer than this recommended time; if complexes are left to sit for 20 mins or longer, transfection efficiency will be drastically reduced (>50% reduction in baculovirus titer produced).

Yes. The ExpiFectamine Sf Transfection Reagent can be used to efficiently transfect Sf9 and Sf21 cells. The reagent can be used to transfect these cells in both adherent and suspension culture formats.

Protein yield can vary greatly from protein to protein. We strongly recommend using the pFastBac 1 - Gus positive control vector (included in the ExpiSf Expression System Starter Kit as well as sold separately - Cat. No. 10360014) to generate Gus-expressing baculovirus and express Gus recombinant protein. The typical yield of Gus protein using the ExpiSf Expression System is approximately 250,000 activity units/mL following the recommended quantification protocol described in the ExpiSf Expression System User Guide. Quantification of the Gus positive control protein will help determine if the low protein yield is due to a low expressing protein, low baculovirus titer/MOI used, poor baculovirus stock quality, a problem with the system components, or transfection and cell culturing conditions. If the expected protein yield is not being achieved with the Gus positive control, we recommend checking the following:

1. Ensure that you have a healthy ExpiSf9 cell culture:
- Cells are >90% viable during normal passaging and at the time of transfection and infection.
- Cells exhibit a doubling time of approximately 24 hrs.
- Cells recover rapidly post-thaw (i.e., reach a cell density of 5 x 10E6 cells/mL with ≥80% viability within 4-5 days post thaw); if not, verify that the culturing guidelines provided in the product manual are being followed, and thaw a new vial of cells if necessary.
- Verify that the incubator temperature is set at 27 degrees C for your cultures as temperatures higher than 28 degrees C may cause cells to overgrow and display reduced yield capacity overtime. Verify that the shake speed is 125±5 rpm for shakers with a 19 mm or 25 mm orbital diameter and 95±5 rpm for shakers with a 50 mm orbital diameter. All of our shake speed recommendations are provided for Nalgene PETG non-baffled Erlenmeyer flasks; if a different culture vessel is being used, additional shake speed and/or other culture parameter(s) optimization may be required.
2. Ensure that you have a high-quality baculovirus stock:
a. Ensure that pure and high-quality bacmid DNA is used:
- Check the quality of the recombinant bacmid by agarose gel electrophoresis to confirm DNA integrity.
- Ensure that a single white colony is picked during bacmid preparation to avoid a mixture of recombinant and non-recombinant baculovirus.
- Confirm that pure bacmid DNA is used (i.e., contains only recombinant bacmid and no empty bacmid), consider screening other DH10Bac transformants (i.e., white colonies).
- Look for the typical signs of a good transfection reaction: When ExpiSf9 Cells are efficiently transfected and baculovirus is being produced, viable cell density is between 80-90% at Day 3 post-transfection and will decrease to <80% at Day 4 post-transfection.
b. Ensure that the recombinant baculovirus contains the gene of interest:
- Verify transposition of bacmid DNA by PCR analysis using the pUC/M13 Forward and Reverse primers.
- Re-transfect ExpiSf9 Cells with new recombinant bacmid DNA, if necessary.
c. We recommend using the suspension-based transfection protocol for generation of high-quality P0 virus, as multiple rounds of viral amplification following the classical adherent format can result in spontaneous excision of the gene of interest as well as the formation of defective viral particles.
d. Ensure that the baculovirus stock is stored properly: Baculovirus stock should be stored at 4 degrees C, protected from light for up to 12 weeks. Alternatively, baculovirus stocks can be frozen and stored at -80 degrees C or in liquid nitrogen (no DMSO or cryopreservative) for longer periods. We do not recommend repeated freeze/thaw cycles of your virus stock. Frozen virus should be stored in small aliquots and not re-frozen once thawed.
3. Ensure that the cells are seeded at 5 x 10E6 viable cells/mL at the time of ExpiSf Enhancer addition.
4. Ensure that the correct amount of ExpiSf Enhancer is added 18-24 hrs prior to infection. Incubating enhancer-treated cells for longer than 24 hrs may result in decreased infection efficiency and low protein yields.
5. Ensure that the cell density at the time of virus infection is at 5-7 x 10E6 viable cells/mL. Infecting cells at higher densities will lead to sub-optimal infection conditions and poor protein yields.
6. Ensure that the optimal volume of baculovirus stock is used to infect cells. We recommend starting with an MOI of 5 and further optimizing the MOI (from 3-10), if necessary.
7. If baculovirus titer wasn't determined, we recommend testing a series of virus stock dilutions to determine the optimal volume to use for virus infections. A starting dilution range of 1:50 - 1:100 (virus stock : culture volume) is a good starting point.