Noncytotoxic DsRed Derivatives for Whole-Cell Labeling Protocol

Introduction of Noncytotoxic DsRed Derivatives for Whole-Cell Labeling

Since the introduction of the green fluorescent protein (GFP) from Aequorea victoria, fluorescent proteins (FPs) have become invaluable research tools for microscopy and flow cytometry. Dozens of FPs have been discovered or engineered to span the visible spectrum. In particular, a number of red FPs have been engineered for use alone or in multi-color studies with GFP. A major goal has been to engineer monomeric red FPs from proteins that are dimeric or tetrameric in their wild-type forms. These efforts have produced the widely-used mFruits as well as DsRed-Monomer, TagRFP, TagRFP-T, mKate, mKate2, mRuby, mKO, and mKO2.

Although monomeric red FPs are theoretically useful for any application, they tend to be dimmer and more photolabile than the corresponding oligomers. Therefore, researchers have also created improved oligomeric red FPs such as DsRed-Express, Kusabira-Orange (KO), TurboRFP, Katushka, and RFP611, RFP637, and RFP639. These oligomeric FPs cannot be used as fusion tags, but they are suitable for labeling organelles, whole cells, tissues, and entire organisms.

A serious problem with both monomeric and oligomeric FPs is cytotoxicity. Most of the available red FPs show pronounced cytotoxicity that presumably stems from aggregation. For this reason, we modified the surface of DsRed Express to create a tetrameric derivative that shows minimal higher-order aggregation. The resulting FP, DsRed-Express2, combines low cytotoxicity with favorable photophysical properties such as brightness, fast maturation, photostability, and pH stability. As a result, the tetrameric DsRed-Express2 is an ideal red FP for whole-cell labeling.

Noncytotoxic color variants were then engineered by modifying the interior of DsRed-Express2. This approach yielded three tetrameric derivatives termed E2-Orange, E2-Red/ Green, and E2-Crimson. Like the parental DsRed Express2, these new variants substantially outperform other FPs with regard to cytotoxicity in bacterial and mammalian systems. E2-Orange is ideal for multi-color experiments involving green and far-red FPs, while E2-Red/Green is useful as a “third color” for flow cytometry in combination with red and green FPs. E2-Crimson is particularly noteworthy because it is the fastest-maturing red or far-red FP, the brightest far-red FP, and the only known member of the GFP family that is efficiently excited with standard 633-nm lasers. As a result, E2-Crimson is useful for multi-color microscopy and flow cytometry. The methods used to engineer and evaluate DsRed-Express2 and its derivatives are detailed below, and can be used as a guide for the development of new noncytotoxic FPs.

Methods of Noncytotoxic DsRed Derivatives for Whole-Cell Labeling

The engineering of useful FPs for whole-cell labeling typically involves multiple rounds of mutagenesis. To generate DsRed-Express2, we employed targeted or random mutagenesis followed by screening for improved variants. Each screen was conducted in two stages, in which a preliminary selection for bright clones was followed by a bacterial-lysis assay to measure aggregation. This general approach can be used to create noncytotoxic derivatives of FPs that have desirable photophysical properties. Oligomeric FPs are best suited to optimization by this strategy, because the multivalent nature of an oligomeric FP enhances the functional affinity that can be detected using the bacterial lysis assay.

In our experience, high solubility of an oligomeric FP in the bacterial-lysis assay correlates strongly with low cytotoxicity in bacterial and mammalian cells. However, cytotoxicity should be measured for each FP. To test for cytotoxicity in bacteria, E. coli colonies expressing a given FP are compared to colonies grown under repressing conditions. Smaller colonies indicate greater cytotoxicity. To test for cytotoxicity in mammalian cells, FPs are expressed in HeLa cells either transiently or by lentiviral transduction, and the fluorescence signal is monitored for several days using flow cytometry. With a cytotoxic FP, the average brightness of the cell population decreases over time, whereas with a noncytotoxic FP, such as enhanced GFP (EGFP) or DsRed-Express2, the average brightness remains steady over time.

Screening E. coli Colonies for Brightness

1. Transform pQE60-NA-based libraries into DH5a cells using standard protocols. Spread the transformation mixture on LB+ampicillin plates at a density that yields approximately 1,000–1,500 colonies per 100-mm Petri plate.
2. After 12–15 h growth at 37°C (see Note 3), examine colony fluorescence using the slide projector, as follows. In a dark room, place an appropriate bandpass filter over the slide projector light source. Wearing an appropriate emission filter goggles, illuminate the Petri plate with the lid off and look for the brightest colonies. Rotate the Petri plate to assure that all regions are examined.
3. Using sterile toothpicks, pick the brightest colonies (approximately 10% of the total) into 96-well plates for screening in the aggregation assay, or restreak promising colonies for maintenance (see Note 4).

Screening for Aggregation Using the Bacterial Lysis Assay

1. Fill each well of a 96-well round-bottom plate with 175 mL of LB+ampicillin. In each plate, set aside four control wells for clones expressing the FP that was the template for mutagenesis (see Note 5). Grow 10–12 h at 37°C (see Note 6), shaking at an angle in an orbital shaker.
2. To store clones for later recovery, use a multichannel pipette to spot 2 mL each from 48 wells onto an LB+ampicillin plate in a grid pattern. Label the Petri plates according to well number. Incubate the Petri plates at 37°C for 12–15 h, and store them at 4°C.
3. Meanwhile, for each 96-well plate, centrifuge the remaining culture samples for 5 min at 3,000×g. Carefully remove the supernatants using a multichannel pipette.
4. Resuspend each cell pellet in 100 mL BPER II using a multichannel pipette. Allow the lysis to proceed for 20 min at 37°C in an orbital shaker.
5. Centrifuge each 96-well plate for 5 min at 3,000×g. Using a multichannel pipette transfer the supernatants to a pair of black 96-well plates, skipping every other column to leave room for the corresponding pellet fractions. Be careful not to create bubbles during the transfer.
6. Using a multichannel pipette, thoroughly resuspend each pellet of lysed cells in 100 mL BPER II. Transfer the resuspended pellet samples to black 96-well plates, placing each pellet sample next to the corresponding supernatant sample.
7. Quantify the fluorescence signals using a 96-well plate fluorometer. It is important to use a fixed gain that is high enough to produce strong signals without overloading the detector. For red FPs, we typically use 560±20 nm excitation and 600±20 nm emission.
8. Determine the aggregation value for each clone by calculating the percentage of the signal found in the pellet. For a given well, divide the pellet signal by the sum of the pellet and supernatant signals. As a positive control, EGFP should have an aggregation value of 2–4%.
9. For each well, compare the aggregation value to the average aggregation value for the four template wells. Calculate the standard error of the mean (s.e.m.) for the four template wells (see Note 7). Multiply the s.e.m. by two and subtract it from the mean aggregation value for the remaining 92 wells, thereby obtaining a threshold for significance. Any well with an aggregation value less than or equal to the threshold should be considered promising for further analysis.
10. To evaluate promising candidates, restreak from the replica spots (step 2) on fresh LB+ampicillin plates. For each candidate, pick four to eight restreaked colonies into 96-well plates, and repeat the aggregation assay with a template control. Candidates that are reproducibly more soluble than the template should be characterized further by preparing miniprep DNA for sequencing.
11. After obtaining the sequences of the improved variants, build a combinatorial library that includes the solubility-enhancing mutations as well as the original codons. In some cases, it may be relevant to test additional mutations at key positions (see Note 8).
12. Screen this library as described above (see Note 9) to identify the optimal combination of mutations.

Assaying Cytotoxicity in Bacteria

1. Adjust the concentrations of pQE-60NA-based FP expression plasmids to 1 ng/mL in sterile H₂O. Transform these plasmids into competent DH10B cells harboring the pREP4 repressor plasmid.
2. Prepare LB+ampicillin+kanamycin plates, one without IPTG and one with 1 mM IPTG. For each plate, use a marker to demarcate sectors for the FPs that are being tested.
3. Spread equal volumes of a given transformation mixture on the appropriate sector of each plate, aiming to achieve a density of about 10 colonies/cm2 (see Note 10).
4. Incubate for 12–15 h at 37°C. Compare colony sizes for repressing versus derepressing conditions. Large colonies under derepressing conditions indicate low cytotoxicity (see Note 11).
5. Colony size is a meaningful measure only if different FPs are expressed at similar levels (see Note 12). Expression is analyzed by SDS–PAGE of cell lysates. For this purpose, grow transformed clones overnight at 37°C in 5-mL cultures of LB+ampicillin+kanamycin to generate saturated precultures. Dilute each preculture to an OD₆₀₀ of 0.05 in 5 mL of LB+ampicillin+kanamycin in a 50-mL baffled flask. After growth for 2 h at 37°C in an orbital shaker to yield an OD₆₀₀ of ~0.5, add 1 mM IPTG to induce FP expression, and incubate with shaking for 4 h at 37°C. Then centrifuge 1 OD₆₀₀ unit of each culture in a microfuge tube, resuspend each cell pellet in 200 mL of 1× SDS–PAGE sample loading buffer, and boil for 10 min. Centrifuge this mixture at 16,000×g for 10 min. Without disturbing the pellet, dilute a portion of each sample supernatant 1:4 in SDS–PAGE sample buffer, and load 20 mL in a lane of an SDS–polyacrylamide gel. Load 10 mL of PageRuler prestained protein ladder as molecular weight standards. Run the gel and stain with Coomassie Blue or a comparable dye. FP bands can be identified based on size and intensity, with reference to a control sample from cells carrying an empty expression vector.

HeLa Cell Transient Transfection Assay for Cytotoxicity

1. Culture HeLa cells in a 100-mm dish to 50–70% confluence.
2. Remove the culture medium by aspiration. Wash the cells with 10 mL sterile PBS. Remove the PBS, and dissociate the HeLa cells by incubating for 4 min at 37°C with 2 mL of Trypsin–EDTA solution. Resuspend the cells in 5 mL of culture medium at 37°C (see Note 13).
3. Add the cell suspension to a 15-mL conical tube and centrifuge for 5 min at 2,000×g.
4. Discard the supernatant, and resuspend the cell pellet in 5 mL of fresh culture medium.
5. Determine the cell density using a hemocytometer.
6. Add 0.5–1×10⁵ cells to each well of several 24-well plates. To test each FP in triplicate, use 15 culture wells per FP plus 5 culture wells for the “No DNA” control. Allow the cells to grow for 16–24 h to attain ~30% confluence.
7. Dilute the FP expression plasmids to 400 ng/mL.
8. For each FP, dilute 30 mL of plasmid in 750 mL of OptiMEM. Mix gently. For the “No DNA” control, use 30 mL of sterile H₂O.
9. In separate tubes, add 15 mL of Lipofectamine 2000 to 750 mL of OptiMEM for each FP. Mix gently and incubate at room temperature for 5 min.
10. For each FP, combine the plasmid mixture with the Lipofectamine mixture. Mix gently and incubate at room temperature for 20 min.
11. For a given FP, add 100 mL of the plasmid/Lipofectamine mixture to each of 15 culture wells. For the “No DNA” control, add 100 mL of the H₂O/Lipofectamine mixture to each of five culture wells.
12. Incubate the cells for 24 h. Remove the Lipofectamine-containing medium and replace with fresh culture medium.
13. Analyze the cells by flow cytometry at daily intervals for 5 days, beginning at 24 h post-transfection. To prepare cells for analysis on a given day, select three wells per FP and one well for the “No DNA” control. For each well, remove the medium, wash the cells with 1 mL of PBS, then replace the PBS with 200 mL of Trypsin–EDTA and incubate at 37°C for 4 min. Remove the Trypsin–EDTA, and suspend the cells in 2 mL of culture medium.
14. Perform flow cytometry with the suspended cells using suitable laser lines and emission filters. For red FPs, 561-nm lasers and PE filter sets (585/15 nm) are commonly used. Based on side scatter area (SSC-A) and forward scatter area (FSC-A) profiles, gate on living cells. Based on side scatter width (SSC-W) and forward scatter width (FSC-W), gate on single, non-aggregated cells. Gate fluorescent cells based on the “No DNA” control sample.
15. Analyze the fluorescence data with FlowJo software. For each sample, including the “No DNA” control, record the average population brightness of living fluorescent cells. To determine the signal/background ratio, divide the average brightness of the fluorescent cells by the average brightness of the “No DNA” cells.

Generation of Lentiviral Vectors and HeLa Cell Lentiviral Transduction Assay for Cytotoxicity

1. Generate lentiviral particles for FP-encoding vectors according to the Lenti-X Expression Kit protocol. In parallel, generate control viral particles by transfecting 293-T cells with pLVX-Puro.
2. Aliquot the viral particles and store them at −80°C (see Note 14).
3. Determine the viral titer for each sample using the QuickTiter Lentivirus Quantitation Kit (see Note 15).
4. Culture HeLa cells to 50–70% confluence. Split the cells into 6-well dishes. Add 2.5×10⁵ cells per well, with three wells per FP and one well for the control virus. Incubate for 16–24 h at 37°C until the cells are ~50% confluent.
5. Remove the culture medium and replace it with exactly 2 mL of fresh culture medium per well.
6. Add 1.6 mL of 10 mg/mL polybrene to each well for a final concentration of 8 mg/mL (see Note 16).
7. Add equivalent titers of viral particles to each well, resulting in three replicates per FP. Incubate for 24 h at 37°C.
8. Remove the culture medium and excess viral particles, wash each well with 2 mL of culture medium, and add 2 mL of fresh culture medium. This time point is defined as day 0.
9. Incubate for 72 h at 37°C to allow expression of the FPs.
10. At the end of this incubation (on day 3), remove the culture medium and wash the cells in each well with 2 mL of PBS. Add 0.5 mL of Trypsin–EDTA to each well, and incubate for 4 min at 37°C.
11. Remove the Trypsin–EDTA, and resuspend the cells in 2 mL of fresh culture medium per well. To allow continued growth, add 0.5 mL of cell suspension to 2 mL of culture medium per well in fresh 6-well plates.
12. Analyze the remaining cells by flow cytometry. Record the average brightness of viable fluorescent cells and control cells. Also record the percentage of viable cells that are fluorescent.
13. Repeat the passage and flow cytometry analysis on days 5 and 7. Do a final analysis on day 10.

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