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Staining cells for FACS at 4 degrees or ambient temperature

Staining cells for FACS at 4 degrees or ambient temperature


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I'm sorting a 293 derived line. One thing that is worrisome for me is cell viability after the sort. Usually I have been staining and washing at room temperature (on the benchtop) on a nutator. I have been reading that most sorting procedures indicate that you should be doing this at four degrees in the dark. It is my understanding that this helps with clumping, but is this at the expense of viability? Does the cold cause these cells to apoptose?

Currently my procedure is the following -

Using HBSS + 1 mM EDTA + 1% BSA (sort buffer) I suspend approx 10-15 million cells per/mL.

  • Stain with primary

  • Spin at 300 x g for 5 minutes

  • Add 15mL of sort buffer, spin again

  • Stain with secondary in 1 mL of sort buffer

  • Wash x2 with 15mL

  • Strain

  • Sort

One thing I can do to minimize cell death is do a live/dead stain (we just added that capability). The other thing I read was to use 50% FBS in your collection tube.

The other thing that might be of concern is that if we are doing many sorts, then some of the tubes may be sitting in sorting buffer for a long time in the queue. How long is too long?

Lastly, we use puromycin and blasticidin as selection markers for our libraries and we usually add these right to the sort after we replate. Should we wait a while?

Thanks for the advice


You have several questions, please try to limit your posts to one question each. That being said, I'll try to address each of your concerns.

Does the cold cause these cells to apoptose?

No. The purpose of staining at 4°C and washing with cold buffer is twofold - to maintain viability (all cellular processes slow down at lower temps) and to prevent receptor internalization. This second point is especially critical as you're using a primary and conjugated secondary, which I really wouldn't recommend for sorting unless absolutely necessary, for example if the antibody you're using isn't available in a directly-conjugated format.

One thing I can do to minimize cell death is do a live/dead stain (we just added that capability).

Live/dead stains are generally a good idea, but make sure the stain you're using isn't cytotoxic. L/D should always be the last staining step before running your samples. A better way (if your sorter supports it) is to use forward scatter (FSC) and side scatter (SSC) height and area measurements to discriminate single, appropriately-sized cells, then gate on them first before gating on your antibody stain (FITC or PE or whatever your secondary is).

The other thing I read was to use 50% FBS in your collection tube.

I'm not sure how much that will help, especially 293s, which are pretty tough. Just make sure you're diluting the collected cells enough in medium to bring the serum percentage back down to 10%.

if we are doing many sorts, then some of the tubes may be sitting in sorting buffer for a long time in the queue. How long is too long?

As long as your samples are kept in the dark on ice or at 4°C, they should be fine for several hours at least (I wouldn't go overnight, though). Also, make sure your collection medium is chilled, as it can shock the cells to go from 4° to 37° too rapidly. Collect your sample, plate it, then put it at room temp or in the incubator (depending on your workflow) and let it warm up that way.

Lastly, we use puromycin and blasticidin as selection markers for our libraries and we usually add these right to the sort after we replate. Should we wait a while?

Nah, don't worry about it. You can wait if you want, or add it right away - I would lean towards adding it more quickly than not, so you don't give unwanted cells any chance of growing out.


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Technology

Download the Certificate of Analysis

Product shipping, storage, shelf life, & solubility

Bioscience kits
The guaranteed shelf life from date of receipt for bioscience kits is listed on the product information sheet. Some kits have an expiration date printed on the kit box label, this is the guaranteed shelf life date calculated from the day that the product shipped from our facility. Kits often are functional for significantly longer than the guaranteed shelf life. If you have an older kit in storage that you wish to use, we recommend performing a small scale positive control experiment to confirm that the kit still works for your application before processing a large number of samples or precious samples.

Antibodies and other conjugates
The guaranteed shelf life from date of receipt for antibodies and conjugates is listed on the product information sheet. Antibodies and other conjugates often are functional for significantly longer than the guaranteed shelf life. If you have an older conjugate in storage that you wish to use, we recommend performing a small scale positive control experiment to confirm that the product still works for your application before processing a large number of samples or precious samples.

For lyophilized antibodies, we recommend reconstituting the antibody with glycerol and antimicrobial preservative like sodium azide for the longest shelf life (note that sodium azide is not compatible with HRP-conjugates).

Chemicals, dyes, and gel stains
Biotium guarantees the stability of chemicals, dyes, and gel stains for at least a year from the date you receive the product. However, the majority of these products are highly stable for many years, as long as they are stored as recommended. Storage conditions can be found on the product information sheet or product safety and data sheet, material safety data sheet, and on the product label. Fluorescent compounds should be protected from light for long term storage.

If you have a Biotium compound that has been in storage for longer than one year that you wish to use, we recommend performing a small scale positive control experiment to confirm that the compound still works for your application before processing a large number of samples or precious samples.

Expiration date based on date of manufacture (DOM)
If your institution requires you to document expiration date based on date of manufacture for reagents, please contact [email protected] for assistance.

Chemical products with special stability considerations:

Ester compounds include the following:
• Succinimidyl esters (SE, also known as NHS esters), such as our amine-reactive dyes
• Acetoxymethyl esters (AM esters) such as our membrane-permeable ion indicator dyes
• Diacetate-modified dyes, like ViaFluor™ 405, CFDA, and CFDA-SE cell viability/cell proliferation dyes

Ester dyes are stable in solid form as long as they are protected from light and moisture. Esters are not stable in aqueous solution. Concentrated stock solutions should be prepared in anhydrous DMSO (see Biotium catalog no. 90082). Stock solutions in anhydrous DMSO can be stored desiccated at -20°C for one month or longer. Esters should be diluted in aqueous solution immediately before use. Succinimidyl esters (SE) should be dissolved in a solution that is free of amine-containing compounds like Tris, glycine, or protein, which will react with the SE functional group. AM esters and diacetate compounds should be dissolved in a solution that is free of serum, because serum could contain esterases that would hydrolyze the compound.

A note on CF™ dye succinimidyl ester stability
Succinimidyl esters are generally susceptible to hydrolysis, which can result in lower labeling efficiency. Heavily sulfonated dyes, such as the Alexa Fluor® dyes, DyLight® dyes and IRDyes® are particularly hygroscopic, worsening the hydrolysis problem. For example, the percent of active Alexa Fluor® 488 succinimidyl ester (SE) could be well below 50% by the time of application (according to the manufacturer’s product datasheet). In a number of Alexa Fluor® SE reactive dyes, the SE group is derived from an aromatic carboxylic acid, while in all of Biotium’s CF™ dyes the SE group is prepared from an aliphatic carboxylic acid. This structural difference reduces the susceptibility of CF™ dye SE reactive groups to hydrolysis, resulting in relatively stable reactive dyes with consistently higher labeling efficiency compared to other SE derivatives of other fluorescent dyes.

Maleimides, MTS and thiosulfate dyes
Like the succinimidyl ester dyes, these dyes are also susceptible to hydrolysis, although generally to a much lower degree. Thus, for long term storage, anhydrous DMSO is recommended for making stock solutions.

Other reactive dyes
Amines, aminooxy (also known as oxylamine), hydrazide, azide, alkyne, BCN, and tyramide reactive dyes, as well as dye free acids, are generally stable in aqueous solution when stored at -20°C for 6-12 months or longer, as long as no compounds are present that may react with the dye’s functional group. See the product information sheets for specific reactive dyes more information.

Coelenterazines and D-luciferin

Coelenterazines are stable in solid form when stored as recommended they are not stable in aqueous solution. Concentrated coelenterazine stock solutions (typically 1-100 mg/mL) should be prepared in ethanol or methanol do not use DMSO or DMF to dissolve coelenterazines, because these solvents will oxidize the compounds. Ethanol or methanol stocks of coelenterazine can be stored at -20°C or below for six months or longer alcohol stocks may evaporate during storage, so use tightly sealing screw cap vials and wrap the vials with Parafilm for long term storage. Propylene glycol also can be used as a solvent to minimize evaporation. If the solvent evaporates, the coelenterazine will still be present in the vial, so note the volume in the vial prior to storage so that you can adjust the solvent volume to correct for evaporation if needed. Prepare working solutions in aqueous buffers immediately before use. Coelenterazines are stable for up to five hours in aqueous solution.

Aquaphile™ coelenterazines are water soluble formulations of coelenterazines. They are stable in solid form when stored as recommended. Aquaphile™ coelenterazines should be dissolved in aqueous solution immediately before use. They are stable for up to five hours in aqueous solution.

Note that coelenterazines are predominantly yellow solids, but may contain dark red or brown flecks. This does not affect product stability or performance. If your coelenterazine is uniformly brown, then it is oxidized and needs to be replaced.

D-luciferin is stable in solid form and as a concentrated stock solution when stored as recommended it is not stable at dilute working concentrations in aqueous solution. Prepare concentrated D-luciferin stock solutions (typically 1-100 mg/mL) in water, and store in aliquots at -20°C or below for six months or longer. Prepare working solutions immediately before use.

Most of our products are stable at room temperature for many days, but we recommend storage at 4°C or -20°C to prolong shelf life. In the case of many of our aqueous dye solutions, the compounds are very stable at room temperature, but we recommend cold storage to prevent the growth of mold or other microbes over time. Therefore, to save on shipping costs, products with recommended storage at 4°C or -20°C may ship at ambient temperature or with an ice pack. These products may thaw without affecting product performance. When you receive the product, place it under the recommended storage conditions.

Most of our products are stable at room temperature for many days, so in all likelihood the product will still work just fine. To be on the safe side, we recommend performing a small scale positive control experiment to confirm that the product still works for your application before processing a large number of samples or precious samples.

One exception that we are aware of is GelGreen™, which is more sensitive to light exposure than most of our other fluorescent dyes. If GelGreen™ is exposed to ambient light for a prolonged period of time (days to weeks), its color will change from dark orange to brick red. If this occurs, the GelGreen will no longer work for gel staining.


Results

Blood refrigeration prior to processing compromises viability and alters composition of recovered PBMC subsets

We initially set out to assess the effect of a delay in processing for refrigerated blood. Samples were processed after storage at 4ଌ for 6, 12 or 24 h, or freshly processed (within 2 h of blood draw). All PBMCs were cryopreserved at �ଌ, then thawed for fluorescence flow cytometric analysis ( Figure 1 ). The frequency of viable cells as determined by NIR viability dye staining decreased with storage at 4ଌ, with significant differences between the control and both the 6- and 24-h groups ( Figure 1 A). Individual cell populations showed differential effects of cold storage CD3 + CD4 + T cells were decreased, CD3 − lymphocytes (B and NK cells) were increased, and CD3 + CD4 − T cells were unchanged. There was also a non-significant trend towards a reduction in the proportion of monocytes, determined as CD4 lo CD14 + ( Figure 1 B𠄾). These data confirm that when stored under refrigerated conditions, a delay in time-to-processing can significantly impact the PBMC populations present in cryopreserved samples.

A total of 4 × 5 ml EDTA blood tubes were collected from each of five donors. Tubes were processed immediately or refrigerated for 6, 12 or 24 h before processing. All PBMCs were cryopreserved and subsequently thawed and assessed by fluorescence flow cytometry for FSC and SSC to distinguish monocytes and lymphocytes, and expression of CD3, CD4 and CD14. (A) Viability was determined using NIR viability dye, with representative dot plots showing changes in forward scatter vs viability over time. Within viable cells, (B) monocytes were gated as CD4 lo CD14 + and their identity confirmed by FSC/SSC, (C) B and NK cells were gated collectively as CD3-negative lymphocytes, (D) CD4 + T cells were gated as CD3 + CD4 + and (E) CD8 + T cells were identified in this limited panel as CD3 + CD4 − . Data were analysed with a one-way repeated measures ANOVA with post-hoc Tukey’s tests and P values indicating significant differences between the timepoints are shown.

Cell recovery is reduced when blood is refrigerated prior to processing

We next sought to understand which stages of the blood processing protocol contributed to these changes in viability and subset recovery, and whether these changes were also evident in samples stored at RT prior to processing. Cell recovery was determined by comparing the number of WBCs recovered from Ficoll-hypaque gradient separation with the number loaded on to the gradient, as determined from the pre-gradient WBC count. At RT, only a small decrease in pre-Ficoll WBC counts was seen (12% over 24 h), a loss of 25% was evident with 24 h of cold storage. After processing, a much greater loss of WBCs was seen for refrigeration times of 6 h (62%) and 24 h (74%), while a small increase for samples stored at RT did not reach statistical significance ( Figure 2 A). In response to refrigeration, we observed the formation of many cell clumps containing erythrocytes, which were not retained at the density interface during PBMC isolation (not shown), and this may account for the poor WBC recovery. Lymphocyte recovery showed the same trends ( Figure 2 B). These observations demonstrate that refrigeration adversely reduced WBC recovery.

A total of 5 × 7 ml heparin blood tubes were collected from each of seven donors, and 2 × 7 ml heparin blood tubes from an additional donor. Individual tubes were either processed immediately or kept at RT or 4ଌ for 6 or 24 h. PBMCs and an aliquot of whole blood were analysed using a Sysmex XP-300™ Automated Hematology Analyzer. These full blood counts were analysed using a one-way mixed effects ANOVA with post-hoc Tukey’s tests. A small drop in WBC was seen with RT storage (5% at 6 h, 11% at 24 h, P=0.04) and a larger drop at 4ଌ (12% at 6 h P=0.003, 25% at 24 h P=0.002). For the additional donor, only the 0 and 24 h RT analyses were performed. Cell recovery of (A) WBC and (B) lymphocytes was calculated as a percentage of the cell number loaded on to the Ficoll-hypaque gradient. Missing values for lymphocytes are due to the inability of the Sysmex to resolve a distinct lymphocyte peak. Data were analysed using a one-way mixed effects ANOVA with post-hoc Tukey’s tests and P values indicating significant differences between the timepoints are shown. For the 4ଌ analysis, a 24-h lymphocyte count was obtained for only a single sample, so the 0- and 6-h timepoints were analysed using a paired values t test.

RT storage of whole blood leads to neutrophil contamination of PBMCs but no changes in major cell subset distribution

Since WBC recovery was markedly reduced by refrigeration, we restricted further analysis to the effects of RT storage. We used fluorescent flow cytometry to assess changes in cellular composition of whole blood and PBMCs after 6 and 24 h RT. There was a slight decrease in the proportion of neutrophils in whole blood, while the proportion of neutrophils in PBMCs increased markedly after RT storage ( Figure 3 A,B). Flow cytometric identification of neutrophils by side scatter and expression of CD16 was confirmed histologically by identification of multilobular-nucleated neutrophils in cytocentrifuge images ( Figure 3 C𠄾).

A total of 3 × 5 ml heparin blood tubes were collected from each of five donors, and individual tubes were either processed immediately or kept at RT for 6 or 24 h. Whole blood and PBMCs were analysed by fluorescence flow cytometry, using a cocktail of antibodies to CD3, CD4, CD8, CD56, CD14 and CD16. Dead cells were excluded with NIR viability dye. The percentage of neutrophils, identified by side scatter and expression of CD16, within live cells was calculated for (A) whole blood and (B) PBMCs. Each donor is indicated by a different colour. Data were analysed with a one-way repeated measures ANOVA with post-hoc Tukey’s tests and P values indicating significant differences between the timepoints are shown. (CE) Aliquots of PBMCs were cytospun, stained with modified Giemsa and photographed. Green arrows indicate neutrophils with typical mature nuclear morphology in the 6 and 24 h samples.

For the other major cell populations (monocytes, NK cells, B cells, T cells and their subsets CD4 and CD8 T cells), storage at RT had no significant effect on their representation within the non-neutrophil fraction of either whole blood or PBMCs ( Figure 4 ). Similarly, viability assessment using Zombie NIR revealed no changes resulting from RT storage. In whole blood, all samples were over 99.4% viable, and in PBMCs the lowest viability recorded was 98.6% (data not shown). These data demonstrate that delaying the blood processing for an extended period of time leads to neutrophil contamination in PBMC preparations.

Data from the experiment described in Figure 3 were analysed to determine the percentage of major cell subsets, expressed as a proportion of non-neutrophil WBCs to account for the effect of low density neutrophil contamination in the 6 and 24 h samples. (A) Total CD3 + T cells, (B) CD56 + NK cells, (C) monocytes, gated for side scatter and CD14 expression, (D) CD19 + B cells, (E) CD4 + T cells and (F) CD8 + T cells. One-way repeated measures ANOVA revealed no significant differences between the timepoints for any of the cell subsets.

Implementing mass cytometry for comprehensive immunophenotypic analysis

The results from fluorescence flow cytometric analysis of blood samples stored at RT suggested that a 24-h delay before processing would have minimal impact on the broad immunophenotype. However the type of immune signature that is increasingly being investigated in cancer patients includes a large number of smaller cell subsets that were not examined in the fluorescence analysis. Aliquots of the PBMC samples from the experiments shown in Figures 2-4 were cryopreserved and thawed for time of flight mass cytometric analysis.

Comparison of the major cell populations identified by mass cytometric analysis of cryopreserved PBMCs ( Figure 5 ) confirmed the results from the fluorescent cytometric analysis performed before cyopreservation ( Figure 4 ). Monocytes, T, NK and B cell numbers were all stable, as were subsets of CD4 and CD8 T cells ( Figure 5 ). CyTOF analysis of neutrophils ( Figure 5 A) confirmed the presence of low density neutrophils within PBMCs prepared after RT storage.

Cryopreserved PBMCs from the experiment described in Figure 3 , plus an additional three donors, were thawed in two batches and analysed using mass cytometry. For each donor, the 0, 6 and 24 h samples were barcoded before staining in a single tube to ensure equivalence of staining conditions across timepoints. Data were gated for the populations indicated. Results for (A) neutrophils are expressed as proportion of total cells, while results for the other cell populations (B–O) are expressed as proportion of total non-neutrophil cells. Treg cells were gated as CD25 + CD127 lo CD4 + T cells. Additional T cell subsets were gated for expression of CD45RA, CD45RO and CCR7 into naïve (CD45RA + CD45RO − CCR7 + ), TCM (CD45RA − CD45RO + CCR7 + ), TEM (CD45RA − CD45RO + CCR7 − ) and TEMRA (CD45RA + CD45RO − CCR7 − ) subsets. Each colour represents an individual donor. One-way repeated measures ANOVA revealed no significant differences between the timepoints for any of the cell subsets, apart from neutrophils. (P) Representative tSNE dimensionality reduction of non-neutrophils across time points for a donor with increasing neutrophil contamination over time (donor 3, represented in dark green in (A–O)). Dots represent individual cells and are coloured by populations in (B–O).

More detailed analysis of the expression of individual cell surface proteins as detected by mass cytometry revealed time-dependent reductions in a subset of markers. These included the chemokine receptors CXCR5, CCR6 and CXCR3, and TIGIT. To illustrate these changes, MSIs for indicative cell subsets were calculated ( Figure 6 ). Interestingly, expression of CD247 (TCRζ) was stable, in contrast with the findings in a previous publication [7].

Data from the experiment described in Figure 5 were analysed to determine the effect of RT blood storage on expression of cell surface proteins detected by metal-labelled antibodies. No proteins increased expression at 6 or 24 h. Proteins whose expression decreased significantly over time, as assessed by MSI, are illustrated in (AE). Calculation of MSI for mass cytometry data is useful only when at least 50% of any cell population have detectable signals in that channel. For this reason, cell subsets with relatively high expression of the indicated proteins are shown. For (A𠄼) CXCR5, CCR6 and TIGIT, expression on all cell subsets was reduced, while for CXCR3, there was a disparity between (D) B cells, (E) CD8 + T cells and (F) monocytes. Data were analysed with a one-way repeated measures ANOVA with post-hoc Tukey’s tests and P values indicating significant differences between the timepoints are shown. (G–I) Expression of CD247 showed no significant change. Donors are identified by the same colours as in Figure 5 .

Reduction in chemokine receptor expression correlates with neutrophil number in PBMCs

Neutrophil contamination in PBMCs has previously been linked to changes in T-cell phenotype associated with whole blood storage before processing [7]. To test whether neutrophil contamination correlated with reduction in chemokine receptor expression, the change in expression of CXCR5 and CCR6 over time, expressed as a percentage of the value at time 0, was graphed relative to the percentage of neutrophils within each individual PBMC sample ( Figure 7 ). There was a statistically significant association between the number of neutrophils contaminating the PBMCs and reduction in expression of CXCR5 and CCR6.

The data shown in Figure 6 were analysed as a function of the percentage of low density neutrophils within PBMCs. (A,B) MSIs for CXCR5 and CCR6 expression by B cells in PBMCs prepared after 6 or 24 h at RT storage were expressed as a percentage of values for 0 h samples and graphed against the percentage of low density neutrophils within PBMCs. Donors are identified by the same colours as in Figures 5 and ​ and6. 6 . (C,D) Linear regression analysis of the correlation between increasing neutrophils and decreasing CXCR5 and CCR6.


Technology

Tech Tips

Download the Certificate of Analysis

Product shipping, storage, shelf life, & solubility

Bioscience kits
The guaranteed shelf life from date of receipt for bioscience kits is listed on the product information sheet. Some kits have an expiration date printed on the kit box label, this is the guaranteed shelf life date calculated from the day that the product shipped from our facility. Kits often are functional for significantly longer than the guaranteed shelf life. If you have an older kit in storage that you wish to use, we recommend performing a small scale positive control experiment to confirm that the kit still works for your application before processing a large number of samples or precious samples.

Antibodies and other conjugates
The guaranteed shelf life from date of receipt for antibodies and conjugates is listed on the product information sheet. Antibodies and other conjugates often are functional for significantly longer than the guaranteed shelf life. If you have an older conjugate in storage that you wish to use, we recommend performing a small scale positive control experiment to confirm that the product still works for your application before processing a large number of samples or precious samples.

For lyophilized antibodies, we recommend reconstituting the antibody with glycerol and antimicrobial preservative like sodium azide for the longest shelf life (note that sodium azide is not compatible with HRP-conjugates).

Chemicals, dyes, and gel stains
Biotium guarantees the stability of chemicals, dyes, and gel stains for at least a year from the date you receive the product. However, the majority of these products are highly stable for many years, as long as they are stored as recommended. Storage conditions can be found on the product information sheet or product safety and data sheet, material safety data sheet, and on the product label. Fluorescent compounds should be protected from light for long term storage.

If you have a Biotium compound that has been in storage for longer than one year that you wish to use, we recommend performing a small scale positive control experiment to confirm that the compound still works for your application before processing a large number of samples or precious samples.

Expiration date based on date of manufacture (DOM)
If your institution requires you to document expiration date based on date of manufacture for reagents, please contact [email protected] for assistance.

Chemical products with special stability considerations:

Ester compounds include the following:
• Succinimidyl esters (SE, also known as NHS esters), such as our amine-reactive dyes
• Acetoxymethyl esters (AM esters) such as our membrane-permeable ion indicator dyes
• Diacetate-modified dyes, like ViaFluor™ 405, CFDA, and CFDA-SE cell viability/cell proliferation dyes

Ester dyes are stable in solid form as long as they are protected from light and moisture. Esters are not stable in aqueous solution. Concentrated stock solutions should be prepared in anhydrous DMSO (see Biotium catalog no. 90082). Stock solutions in anhydrous DMSO can be stored desiccated at -20°C for one month or longer. Esters should be diluted in aqueous solution immediately before use. Succinimidyl esters (SE) should be dissolved in a solution that is free of amine-containing compounds like Tris, glycine, or protein, which will react with the SE functional group. AM esters and diacetate compounds should be dissolved in a solution that is free of serum, because serum could contain esterases that would hydrolyze the compound.

A note on CF™ dye succinimidyl ester stability
Succinimidyl esters are generally susceptible to hydrolysis, which can result in lower labeling efficiency. Heavily sulfonated dyes, such as the Alexa Fluor® dyes, DyLight® dyes and IRDyes® are particularly hygroscopic, worsening the hydrolysis problem. For example, the percent of active Alexa Fluor® 488 succinimidyl ester (SE) could be well below 50% by the time of application (according to the manufacturer’s product datasheet). In a number of Alexa Fluor® SE reactive dyes, the SE group is derived from an aromatic carboxylic acid, while in all of Biotium’s CF™ dyes the SE group is prepared from an aliphatic carboxylic acid. This structural difference reduces the susceptibility of CF™ dye SE reactive groups to hydrolysis, resulting in relatively stable reactive dyes with consistently higher labeling efficiency compared to other SE derivatives of other fluorescent dyes.

Maleimides, MTS and thiosulfate dyes
Like the succinimidyl ester dyes, these dyes are also susceptible to hydrolysis, although generally to a much lower degree. Thus, for long term storage, anhydrous DMSO is recommended for making stock solutions.

Other reactive dyes
Amines, aminooxy (also known as oxylamine), hydrazide, azide, alkyne, BCN, and tyramide reactive dyes, as well as dye free acids, are generally stable in aqueous solution when stored at -20°C for 6-12 months or longer, as long as no compounds are present that may react with the dye’s functional group. See the product information sheets for specific reactive dyes more information.

Coelenterazines and D-luciferin

Coelenterazines are stable in solid form when stored as recommended they are not stable in aqueous solution. Concentrated coelenterazine stock solutions (typically 1-100 mg/mL) should be prepared in ethanol or methanol do not use DMSO or DMF to dissolve coelenterazines, because these solvents will oxidize the compounds. Ethanol or methanol stocks of coelenterazine can be stored at -20°C or below for six months or longer alcohol stocks may evaporate during storage, so use tightly sealing screw cap vials and wrap the vials with Parafilm for long term storage. Propylene glycol also can be used as a solvent to minimize evaporation. If the solvent evaporates, the coelenterazine will still be present in the vial, so note the volume in the vial prior to storage so that you can adjust the solvent volume to correct for evaporation if needed. Prepare working solutions in aqueous buffers immediately before use. Coelenterazines are stable for up to five hours in aqueous solution.

Aquaphile™ coelenterazines are water soluble formulations of coelenterazines. They are stable in solid form when stored as recommended. Aquaphile™ coelenterazines should be dissolved in aqueous solution immediately before use. They are stable for up to five hours in aqueous solution.

Note that coelenterazines are predominantly yellow solids, but may contain dark red or brown flecks. This does not affect product stability or performance. If your coelenterazine is uniformly brown, then it is oxidized and needs to be replaced.

D-luciferin is stable in solid form and as a concentrated stock solution when stored as recommended it is not stable at dilute working concentrations in aqueous solution. Prepare concentrated D-luciferin stock solutions (typically 1-100 mg/mL) in water, and store in aliquots at -20°C or below for six months or longer. Prepare working solutions immediately before use.

Most of our products are stable at room temperature for many days, but we recommend storage at 4°C or -20°C to prolong shelf life. In the case of many of our aqueous dye solutions, the compounds are very stable at room temperature, but we recommend cold storage to prevent the growth of mold or other microbes over time. Therefore, to save on shipping costs, products with recommended storage at 4°C or -20°C may ship at ambient temperature or with an ice pack. These products may thaw without affecting product performance. When you receive the product, place it under the recommended storage conditions.

Most of our products are stable at room temperature for many days, so in all likelihood the product will still work just fine. To be on the safe side, we recommend performing a small scale positive control experiment to confirm that the product still works for your application before processing a large number of samples or precious samples.

One exception that we are aware of is GelGreen™, which is more sensitive to light exposure than most of our other fluorescent dyes. If GelGreen™ is exposed to ambient light for a prolonged period of time (days to weeks), its color will change from dark orange to brick red. If this occurs, the GelGreen will no longer work for gel staining.


Introduction

Phytoplankton are responsible for about 50% of Earth’s primary production [1], with diatoms accounting for about half of this [2,3]. As primary producers, phytoplankton form the basis of most marine food webs. In addition, diatoms and phytoplankton in general can serve as indicators of ecosystem change since they are particularly sensitive to shifts in environmental conditions. This is also important in the context of regional and global change, since human activities and associated increases in greenhouse gas emissions have led to simultaneous changes in a number of aquatic abiotic parameters. Consequently, phytoplankton are exposed to the simultaneous effects of multiple anthropogenic stressors, such as increasing temperature and pCO2, as well as shifts in dissolved nutrient concentrations and stoichiometry.

Phytoplankton can respond to changes in environmental conditions at different levels. At the first level, the phenotypic plasticity of individual phytoplankton cells largely determines whether the cells can subsist under changing abiotic conditions. How well a single strain copes with changing environmental conditions depends on the tolerance range of individual cells [4,5]. At the second level, the plasticity of different strains within a population is the critical factor. High plasticity of individual strains has the potential to buffer negative effects of environmental change, while low plasticity of these strains can lead to shifts in population structure [6]. At this level, the sum of the tolerance ranges of different strains within a population determines whether the population can cope with environmental change. The third level of phytoplankton response results in a change in community composition when altered abiotic conditions lead to species selection. Despite the importance of all three levels of response (within strains, between strains, between species), many studies have focused on the response of community [7–9], populations [10,11], and strains [6,12,13], while much less attention has been paid to the phenotypic plasticity of individual cells within a strain.

One of the most important characteristics or traits of individual phytoplankton cells that determine their ecological success is cell size [reviewed by 7], as it is directly correlated with many other traits and is influenced by a range of environmental factors. Smaller cells have lower sinking rates [14], better nutrient uptake due to a higher surface-to-volume ratio, and a smaller diffusive boundary layer [15,16]. On the other hand, smaller cells are often more susceptible to grazing [17]. Thus, cellular characteristics not only affect the ecological role of plankton but are also shaped in part by environmental conditions. Changing temperature and pCO2 regimes could, therefore, have an impact on cellular traits, with consequences for trophic and system processes. So far, most studies of phytoplankton traits and their response to environmental drivers have been conducted using bulk measurements. For example, Hillebrand et al. [18] observed that phytoplankton exhibit large variability in cellular nutrient content at low growth rates and that cellular nitrogen to phosphorus (N:P) ratios converge with increasing growth rate. Hence, faster growth causes cultures to become more similar and their variability to decrease. This phenomenon was first observed by Goldman in 1986 [19]. However, the study by Hillebrand et al. compared different populations and different species, and although the authors conclude that this should be the case, it remains unclear whether the above-mentioned patterns also apply within individual strains. Given the importance of phenotypic plasticity in coping with altered environmental conditions, a proper evaluation of the response of phytoplankton to global change cannot be made without studying their phenotypic plasticity and cell-to-cell variability within individual strains. Flow cytometry, which measures traits of individual cells, is a well-established method to investigate the variability of cellular characteristics such as cell size, pigments, and biochemical components [20–22].

Since current CO2 concentrations are not saturating for Rubisco, the enzyme that catalyzes primary fixation of inorganic carbon [23], higher pCO2 resulting from anthropogenic emissions can potentially favor photosynthesis and phytoplankton growth [24–26]. Furthermore, rising water temperatures caused by increased atmospheric CO2 can affect physiological processes, including growth, resource acquisition, and photosynthesis [27,28]. Additionally, human activities influence the concentrations and ratios of dissolved nutrients such as nitrogen (N) and phosphorus (P), which are essential for phytoplankton growth. Legal restrictions on nutrient inputs to the environment have resulted in increasing N:P ratios, which are predicted to rise even further in the future, as atmospheric N deposition is likely to continue to increase [29], and P reserves on Earth could be depleted in 50 to 200 years if current levels of use are maintained or increased [30,31]. However, as environmental conditions change, the elemental stoichiometry of phytoplankton species and their biochemical requirements will also be affected [25]. Given the influence of these global change drivers on phytoplankton growth rate and the strong correlation of growth rate with other traits such as cell size [25], it is crucial to investigate the phenotypic plasticity of multiple phytoplankton traits, and how the traits are correlated.

Considering that changes in pCO2, temperature, and nutrient concentrations do not happen separately, but rather simultaneously, we conducted a multi-driver experiment with realistic future conditions based on predictions from the Intergovernmental Panel on Climate Change (IPCC). We tested the RCP 6.0 and 8.5 scenarios by simulating a temperature increase of 1.5 and 3°C combined with increasing pCO2 up to 800 and 1000 ppm, respectively, and we combined these scenarios with an increase in dissolved N:P ratios from 16 to 25. Using a single-cell approach, we analyzed the response of a single strain of the diatom species Thalassiosira weissflogii, currently also named Conticribra weissflogii. Since we acquired the strain we worked with from an algal culture bank under the name T. weissflogii, we will retain the generic name Thalassiosira in this paper. T. weissflogii is mostly found in temperate brackish waters, but it has also been isolated all around the globe in areas ranging from the Long Island Sound, New York, USA, to Jakarta Harbor, Indonesia, to the German Bight (North Sea), as well as in the middle of the Pacific Ocean [32–34] and can, therefore, be considered as a cosmopolitan species. Furthermore, T. weissflogii is known to react in a number of ways to altered abiotic conditions [35,36]. Sugie and Yoshima [37] reported a decrease in cell size of T. weissflogii under increased pCO2 and Wu et al. [38] identified a slight increase of growth rate at higher pCO2. Nevertheless, the single-cell plasticity of multiple traits of T. weissflogii has never been investigated. Since measurements on a single-cell level require certain criteria regarding cell size and chain-formation, we used this well-studied diatom species to analyze the link between growth rate, cell size, and biochemical composition of cells growing under different future global change conditions. More specifically, we tested the hypothesis that not only between species and populations [18] but also within one single strain, cell-to-cell variability of phytoplankton decreases with increasing growth rate. Using flow cytometry, we are able to analyze both, the reaction of single strain populations to different environmental conditions by studying the variation between cultures, as well as the variability within one replicate at a distinct growth rate. The former, we describe in the following as among-strain population variability and the latter as cell-to-cell variability within a single strain population. Both concepts can be used to assess phenotypic plasticity which plays an important role when addressing the question of the effect of global change on phytoplankton.


RESULTS

Calibration of Lactadherin/Annexin V Assay

We hypothesized that the PS-binding properties of lactadherin would enable detection of early PS exposure on dying cells. Preliminary binding studies that related PS content to binding of lactadherin and annexin V were performed with synthetic membranes supported by 2-μm glass microspheres ( 26 ) (Fig. 1). The Ca ++ concentration was 1.5 mM near the upper limit of ionized Ca ++ in blood plasma ( 28 ). Because the rate of PE exposure during early apoptosis has not been measured, we evaluated binding to synthetic membranes with minimal and maximal PE content. First, we assumed that PE would not be exposed together with PS. The PS content was varied from 0 to 12% in synthetic membranes with 2% PE (Fig. 1A). The quantity of lactadherin bound increased with PS content and approached a plateau at 8% PS. In contrast, annexin V binding was not detected until the PS content was 8%, after which it increased rapidly. Second, we assumed that the rate of PE efflux would exceed the rate of PS efflux. Synthetic membranes were synthesized with a PE:PS ratio of 4:1 (Fig. 1B). Again, lactadherin binding was proportional to PS content whereas the relationship between PS and annexin V binding was sigmoidal. The presence of excess PE reduced the PS threshold for annexin V to ∼2.5% PS (Fig. 1B). These results indicate that lactadherin can be expected to bind to cell membranes with PS content ≥0.5%, while annexin V is predicted to bind when the PS content exceeds 2.5%–8%, depending on the PE content. Thus, cell membranes that stain with lactadherin, but not annexin V are likely to have membrane PS content <8% and possibly <2.5%.

Similar results were obtained with annexin V conjugated to three different fluorophores, supplied by three vendors. In addition, a conjugate of annexin V-Alexa Fluor 647 prepared in our laboratory yielded the same pattern of binding. The Alexa Fluor 647 conjugate was utilized for the experiments displayed in Figures 1C and 2–5.

Competition binding experiments were performed to determine whether lactadherin and annexin V could compete with each other for the same binding sites (Figs. 1C and 1D). Unlabeled annexin V competed for only about 40% of the lactadherin binding sites on membranes with 4% PS and 15% PE. By comparison, unlabeled lactadherin competed for ∼70% of annexin V binding sites on these membranes. The results suggest that the binding sites for lactadherin and annexin V overlap but are not identical. They indicate, further, that the presence of either protein at concentrations less than 8 nM would not diminish binding of the other by more than 20%. The arrows indicate the degree of displacement predicted for each protein for experiments depicted in Figures 3–5.

Temporal course of lactadherin binding. Flow cytometry analysis indicated that control HL-60 cells exhibited little staining with lactadherin or PI (upper panel-open arrow). After 24 h of treatment with etoposide, three intensities of lactadherin staining were evident (black arrow, open arrowhead, black arrowhead). After 48 h, more than 95% of cells were positive for both lactadherin and PI. With the displayed quadrant markers, the cells positive for both lactadherin and PI increased from 2.4% for controls to 90.4% at 48 h. Contour lines are drawn at intervals of 15% linear event density. The quantity of lactadherin binding to K-562 cells and HL-60 cells at various time points was compared with overlaid histograms (lower panel). The intensities of staining identified on the contour plot are indicated by the same arrow and arrowhead symbols at the same fluorescence intensities. Most HL-60 cells stained with lactadherin at an intermediate level after 24 h and brightly after 48 h. The mean progression of K-562 cells staining by lactadherin in eight experiments is depicted in the lower right panel. The fluorescence intensity gates (M1–M4) are depicted in the center panel, labeled “K-562”. Results are representative of three experiments (upper panel) five experiments (lower left and center panels).

Lactadherin Binding to Apoptotic Cells

We wished to confirm that lactadherin could detect PS exposure on apoptotic cells. Therefore, K-562 and HL-60 cells were treated with etoposide for 48 h and evaluated with conventional stains as well as lactadherin (Fig. 2).

Staining with hematoxylin and eosin confirmed that some K-562 cells had condensed and fragmented nuclei (Fig. 2A). Positive TUNEL staining confirmed that the DNA was fragmented in these cells (Fig. 2B). Costaining cells with lactadherin and propidium iodide (PI) (Fig. 2C) confirmed that PI permeable cells expose sufficient PS to enable diffuse binding of lactadherin. The diffuse blebs that appear transiently in the course of apoptosis ( 29 ) were identified on a small number of cells (Figs. 2D and 2E). These vesicles stained diffusely with lactadherin. Therefore, K-562 cells expose sufficient PS to bind lactadherin as they undergo programmed cell death.

The engagement of PS-binding motifs in staining by lactadherin and annexin V was evaluated in competition binding experiments (Figs. 2F and 2G). Phospholipid vesicles containing 8% PS competed with etoposide-treated cells for both lactadherin and annexin V binding. These vesicles competed for more than 95% of lactadherin binding but only about 85% of annexin V binding. This difference was rationalized by assuming that a larger fraction of annexin V was internalized by the cells ( 16 ). The results confirm that the PS-binding motifs of both lactadherin and annexin V participate in binding to the etoposide-treated apoptotic cells. Addition of EDTA at twice the Ca ++ concentration reduced staining by annexin V but did not decrease lactadherin binding as detected by flow cytometry or by microscopy (not shown). These results confirmed the Ca ++ -dependent PS binding of annexin V and the Ca ++ -independent PS binding of lactadherin. Furthermore, they exclude the possibility that lactadherin binding was mediated by interaction with the αvβ3 or αvβ5 integrins of the dying cells.

Experiments were conducted to determine whether lactadherin could detect PS exposure before cells exhibit increased permeability to PI (Fig. 3, upper panel). Twenty-four hours after initiation of etoposide exposure, three cell populations were evident, indicating the progressive pathway of PS exposure and PI permeability. Approximately 25% of cells remained in the left lower quadrant (solid arrow), but PS exposure was greater than that of untreated cells (open arrow). Approximately 25% were positive for lactadherin staining but negative for PI (open arrowhead). Another 25% exposed the same quantity of PS but were also stained with PI (upper open arrowhead). Finally, a population of cells stained positively for PI and stained very strongly for lactadherin (solid arrowhead). After 48 h the majority of cells stained maximally with both PI and lactadherin. Samples studied at intermediate time points confirmed that most cells progressed through the two intermediate stages identified after 24 h (not shown). These findings indicate that PS exposure begins prior to PI permeability, but that complete PS exposure is delayed for many hours after PI permeability.

Lactadherin staining of both K-562 and HL-60 cells were monitored over 72 h to determine the comparative rates of PS exposure (Fig. 3, lower panel). The results were consistent with progressive PS exposure after treatment with etoposide for both cell types. The arrows and arrowheads on the HL-60 histogram depict the modal fluorescence intensities for untreated cells as well as minimally, intermediate, and maximally responding populations (Fig. 3, 24 h). The elapsed time between etoposide exposure and PS exposure differed for HL-60 cells vs. K-562 cells. HL-60 cells stained within 6 h of etoposide (not shown) while K-562 cells did not stain until at least 12 h had elapsed since etoposide exposure was initiated. Lactadherin staining was maximal for HL-60 cells by 48 h but required at least 72 h treatment for K-562 cells. The mean progression of PS exposure for eight separate experiments is depicted in the lower right panel. The gates utilized to detect the fraction of cells with each quantity of PS exposure are in the K-562 panel. These findings indicate that PS exposure is progressive over more than 24 h for both K-562 and HL-60 cells.

Costaining with Lactadherin and Annexin V

We utilized flow cytometry to evaluate simultaneous staining by lactadherin and annexin V (Fig. 4, upper panel). Dot plots indicate that most cells proceed through a pathway in which staining by lactadherin is increased to a greater degree than by annexin V. For example, prior to etoposide exposure, only 0.6% of K562 cells stained with lactadherin. After 24 h exposure to etoposide, the fraction of the population in the “lactadherin positive” region increased to 32.6%. After 72 h of etoposide exposure, 84.1% were strongly positive for both annexin V and lactadherin. Comparative staining with lactadherin and PI vs. Annexin V and PI confirmed that the two proteins have distinct staining patterns prior to PI staining (not shown). Lactadherin stained the majority of cells with a continuous intensity from negative to highly positive. In contrast, staining with annexin V showed most cells with either minimal staining or bright staining immediately prior to permeability to PI. Very few cells demonstrated intermediate levels of annexin V staining, indicating that most cells had not exposed sufficient PS to reach the annexin V threshold (Fig. 1). Confocal microscopy revealed the cellular features that display PS exposure after 24 h of etoposide exposure (Fig. 4, lower panels). Many cells had minimally changed gross architecture (black arrow) while a similar number were condensed (arrowhead). The normal-sized cells had patches and small appendages that stained with lactadherin (white arrows), but stained weakly or not at all with annexin V. The condensed cells had diffused lactadherin staining, detected as rings, but annexin V staining was punctate. Most cells exhibited staining of internal cell bodies by annexin V. In fact, the internalized bodies were the primary location of annexin V staining rather than the cell surface. These results indicate that an early response to etoposide treatment is localized PS exposure and that the quantity of PS exposure generally remains below the annexin V threshold.

PS exposure time course and topography detected by costaining with lactadherin and annexin V. PS exposure on K-562 and HL-60 cells was evaluated by costaining with lactadherin and annexin V after 24, 48, and 72 h of etoposide exposure. Flow cytometry dot plots indicated that most cells stain dimly with lactadherin after 24 h and stain less intensely for annexin V. After 72 h nearly all cells stained intensely with both lactadherin and annexin V. The fraction of cells in the quadrants indicating positive staining for both lactadherin and annexin V increased from 0.2% to 84%. Confocal microscopy of cells was performed after cells were treated for 24 h. Phase contrast illustrates cells of normal appearance (arrow) as well as smaller, spherical cells (arrowhead). Annexin V stained internal bodies of spherical cells as well as some discrete membrane features. Lactadherin bound diffusely to the plasma membranes of some shrunken, spherical cells and stained focal and protruding membrane features arrows of normal-size and some shrunken cells. A fluorescence overlay illustrates features where staining by lactadherin and annexin V are coincident (yellow) and features stained preferentially by annexin V (red) and lactadherin (green). In some experiments the Ca ++ concentration was increased to 3 mM. Results are displayed as an overlay of red and green fluorescence in addition to phase contrast (yielding a blue background). Annexin V was found primarily on internal bodies while lactadherin was localized to the plasma membrane. An enlargement of a cell undergoing vesiculation is presented to better illustrate the pattern of lactadherin vs. annexin V localization on these cells. K-562 cells are depicted in upper seven panels, and HL-60 cells in lower two panels. Results representative of at least five experiments.

Annexin V and lactadherin staining studies were also performed with Ca ++ increased from 1.5 to 3 mM (Fig. 4), similar to the conditions generally employed to probe apoptotic cells for PS exposure with annexin V. The results showed an equivalent pattern of staining with lactadherin and annexin V. The blue background reflects superimposition of the phase contrast with fluorescence images. An enlarged view of a vesiculating cell, with 3 mM Ca ++ , highlights the preferential binding of lactadherin to membrane vesicles and the preferential localization of annexin V to internal bodies. Experiments were performed with increasing annexin V concentrations, as other investigators have sometimes utilized concentrations much higher than the apparent dissociation constant of <5 nM ( 30 ). These conditions led to increased staining of both etoposide and control cells to approximately the same degree. The results suggest that use of annexin V for selective staining of exposed PS is probably most effectively performed with annexin V concentrations <6 nM.

After 48 h of exposure to etoposide, the majority of K-562 cells exposed sufficient PS to stain with both lactadherin and annexin V (Fig. 5). Some cells had fragmented nuclei and irregular contours, changes that are consistent with completed apoptosis (Fig. 5A, arrowhead). Most cells were round with condensed, but regular nuclei, consistent with the ongoing cell death program (Fig. 5A, arrows). Most of the condensed cells stained diffusely with lactadherin (Fig. 5C, white arrows). Other cells, with a lesser degree of condensation, had scattered membrane appendages that stained with lactadherin, similar to dominant cell morphology observed at 24 h. Annexin V stained internal bodies of the cells with intact nuclei (Fig. 5B) confirming that significant annexin V is internalized over the 10 min time course of these experiments ( 16 ). Annexin V also stained discrete patches on the plasma membranes of these cells. A composite image (Fig. 5D) contrasts cell features that stain preferentially with lactadherin (green) annexin V (red) and highlights regions that stained with both lactadherin and annexin V (yellow).

Progressive PS exposure of K-562 cells costained with lactadherin and annexin V after 48 h etoposide exposure. Phase contrast microscopy (A) showed cells of normal size and contour (white arrowhead), shrunken, spherical cells (small arrows) and irregular cells with fragmented nuclei (black arrowhead). Annexin V (B) was identified in intracellular bodies (arrowhead) and in discrete surface patches in a large-portion of the cells. Lactadherin (C) stained discrete surface patches and protuberances, diffusely stained the membranes of the shrunken, spherical cells (arrows) and brightly stained most of the irregular cell. An overlay image (D) identifies cell features that co-stain with lactadherin and annexin V yellow as well as areas that stain preferentially with each protein. A higher magnification overlay of different cells (E) shows better the demarcation between internal bodies that stain for annexin V and lactadherin-binding plasma membrane. Contour profiles of lactadherin staining intensity (F) and annexin V staining intensity (G) illustrate the cell features that stain with lactadherin as well as the relative intensity of staining of different cells. Results are representative of at least five experiments.

The contrasting pattern of staining with lactadherin vs. annexin V is highlighted by the profile analysis of costained cells (Figs. 5E–5G). A composite image at higher magnification demonstrates that some internal bodies stain for both lactadherin and annexin V, though predominantly for annexin V. Color intensity profiles of the same cells illustrated that lactadherin diffusely stained the plasma membrane of cells progressing through the cell death program and that the intensity increased with progressive apoptosis (Fig. 5F). Annexin V stained focal regions of the plasma membrane on these cells and internal bodies (Fig. 5G). Thus, these results indicate that dying leukemoid cells expose PS in three stages. PS exposure on most of the cell membrane remains below the annexin V threshold until the final stage of apoptosis.


Freezing and thawing of cells - (Sep/09/2004 )

Though my experience of handling the cells is less than 2years. I would like to suggest you some things. Even though if you leave DMSO in the media after thawing the cells the dmso in low cocentration is not toxic 2 cells more over once you dilute the small volume of cell suspension in 5-10 ml of media, it further dilutes the dmso. It looks to me like the protocol you are following may be causing the cells to die, I think you recognise the fact that centrifuging at high speed can lead to cell necrosis. One method is to lower the speed of centrifuge maybe 900-1000 rpm for 4-5 min should be adiquate.
Alternatively you should also consider the method you are using for preserving the cells. If you freeze the cells directly in liquid nitrogen this can also lead to cold shock induced cell lysis or damage the membrane, one way of preventing this is gradually lowering the temperature such as storing 4 degrees for few hours then transfering to -20 degree for overnight and then storing in -70 degrees for an couple of days then transfering to liquid nitrogen tanks, by following this method u r not only making the cells to adjust to the lower temp you are also ensuring that they donot undergo cold induced necrosis leading to cell membrane damage and lysis.

I hope this information is of use to you.

If you need any further information you can mail me at [email protected]

I dont agree with preethi or anil as one cannot preserve the cells in glycerol for more than 6-8 months.

Why do you want to pellet your cells after thawing. From your procedure you are using DMSO (1:10). DMSO will not give toxicity at low concentration
My suggestion is to add thawed cell suspn to 5-8 ml culture medium and incubate for a day then change medium. Initial pelleting could be avoided.
I never do a pelleting when I thaw cells (from -85 deg C). I never had a problem with low percentage in revial.

I am experiencing similar problems with my LCL b cell line. See topic title:

LCL B cell culture - Death within 24 hours of thawing.

I might try some of the suggestions here.

hi
I am facing problems in thawing of cells. The procedure which we follow is- we take stock from liq. N2 then though it in water bath at 37 degree. Then we take medium in falcon (5ml) then add thawed cells dropwise into it. We then centrifuge it at 1500rpm for 5 mins.
but the problem is that after thawing i am unable to get viable cells.
Almost all cells are dead. So what all modification shall i make ? Its really hard to revive cell line from frozen stock?

This is how to freeze and thaw cells:

Always check the cell viability before freezing. They should be highly viable: about 95%. keep your freezing media ( 10% DMSO in FBS) cold. Centrifuge cells for 1000 RMP for 5 minutes. Prepare labeled cryogenic vials. Cell concentration should be about for e.g. collected from a 50 ml flask, about 50 million cells. You can have 10 vials and about 0.5 milion cells per vial. After you centrifuge, get rid of media completely, gently tap the pellet to make it loose. Add 5 mls of cold freezing media resuspend with a pipet and transfer 0.5ml to each vial. Close the cap tight and place cryogenic tubes in special cryogenic container that has alcohol at the bottom and cool down gradually, they usually hold up to 20 vials. Close the top and transfer the container to -70 freezer. Wait 24 hours and no longer than 15 days before transferring them out of the container into the liquid nitrogen boxes.

To thaw the cells, be very quick, take the viall out still with some liquid nitrogen, walk to the water bath. Take the vial, make sure the cap is very tight, sometimes it becomes loose. thaw the vial holding the opening upward so the water from the water bath does not contaminate cells or if there is some detegent in the water it doesn't become in contact with the inside of the vial . When frozen cell media is almost half thawed (about few minutes), take it under the hood and add 0.5 warm media to the cells and immediately transfer to a flask with about 10 to 15ml warm media in it. DMSO will be diluted and you can check the viability.

Good luck and let us know if it had worked.

When I went through top to bottom of this page, it remind me something I could tell in general

As the wise words (Auguste Rodin) say
"Nothing is a waste of time if you use the experience wisely"

We have -85 Deep Freezer for cell storage in our lab. I got normal revival of cells even after an year, when I preserved them in glycerol. Those were cell lines, not primary cells. The case may be different with primary cells. Yet to get experience with it.

I work with cells not more than half year, but I did met with the similar problem with you. And it's strange that I did all the operation right: thaw quickly, no shaking, and no bubbles when adding medium. But still, I can't get the cell alive, just they don't settle down or very very slow. Plus, I never spin my cell, because I always dilute them into T-75 flask at which concentration DMSO will not be toxic. So the only changes I made I guess is to make sure medium really warm, not just warm them up in 37 degree for 15mins, but 30mins to make sure.

Hi, there
I think the cell line is more tough. I do the similar way to freeze and thaw cells as you suggest above. It works fine. But when I come to fresh spleen cells, I have only 10 percent cells viable after freeze and thaw. My question is that if I am not going to culture the cells after I thaw them, instead I want to do FACS staining and cell sorting, should I dilute them in cold media or warm media after I thaw them in the water bath? I always leave cells in 37 degree water bath for just one min, but they are still crystal. I don't know whether I can leave them in the water bath for a little bit longer. Thank you!

Spleen cells could survive freeze and thawing as well. I would freeze the same number of cells in two separate containers and thaw them in warm or room temperature media to see the difference atfer 24 hours in 37C0 incubator. I usually take the tube out of the liquid nitrogen then warm it up in the water bath for may be one minute and while still is half frozen i add 37degree media to dilute the DMSO. If spleen cells have been treated with amonium chloride for lysing red blood cells they will become very sensitive to freez/thawing procedure.


Product Details

Target Species Human Product Form Purified IgG conjugated to Alexa Fluor® 488 - liquid Product Form Purified IgG conjugated to Alexa Fluor® 647 - liquid Product Form Purified IgG - liquid Product Form Purified IgG conjugated to R. Phycoerythrin (RPE) - lyophilized Reconstitution Reconstitute with 1 ml distilled water Preparation Purified IgG prepared by affinity chromatography on Protein G from tissue culture supernatant Preparation Purified IgG prepared by affinity chromatography on Protein A from tissue culture supernatant Preparation Purified IgG prepared by affinity chromatography on Protein G from tissue culture supernatant Preparation Purified IgG prepared by affinity chromatography on Protein G from tissue culture supernatant. Buffer Solution Phosphate buffered saline Buffer Solution Phosphate buffered saline Buffer Solution Phosphate buffered saline Buffer Solution Phosphate buffered saline Preservative Stabilisers
0.09%Sodium Azide
1%Bovine Serum Albumin
Preservative Stabilisers
0.09%Sodium Azide
1%Bovine Serum Albumin
Preservative Stabilisers
0.09%Sodium Azide
Preservative Stabilisers
0.09%Sodium Azide
1%Bovine Serum Albumin
5%Sucrose
Carrier Free Yes Approx. Protein Concentrations IgG concentration 0.05 mg/ml Approx. Protein Concentrations IgG concentration 0.05 mg/ml Approx. Protein Concentrations IgG concentration 1.0 mg/ml Fusion Partners Spleen cells from immunised BALB/c mice were fused with cells of the P3U1 myeloma cell line.

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