The TokuKit is Teiko’s whole blood collection and preservation kit, designed for immune cytometry profiling in research studies and clinical trials. It simplifies blood sample preservation with a quick, 3-step process—collect, fix, and freeze—that takes 20 minutes or less. No specialized training or equipment is needed.
Samples processed with the TokuKit are frozen at -80°C and remain stable for months, unlike other preservation methods like Transfix or Cyto-Chex, which are limited to days. Unlike PBMC isolation, the TokuKit preserves granulocytes—neutrophils, basophils, and mast cells—enabling a more complete profile of the immune system.
Learn more at teiko.bio/tokukit.
Blood samples processed with Teiko's TokuKits can be sent directly to our CLIA-compliant laboratory in Salt Lake City, Utah. They can be received Monday–Friday, 9 a.m.–5 p.m.
Address:
675 Arapeen Drive, Suite 301
Salt Lake City, UT 84108
ATTN: Li-Chun Cheng or Teiko Lab
We and others (e.g. Rahman, Tordesillas and Berin, Cytometry Part A 2016) have found that eosinophils are difficult to reliably identify by mass cytometry, even when eosinophil-specific markers like SIGLEC-8 are included in the panel. In our validation studies, we observed that eosinophils can bind nonspecifically to certain antibodies for antigens they don’t express. This can result in eosinophils being misidentified as other cell types, such as platelets or neutrophils.
We also found that eosinophil frequencies are highly sensitive to storage conditions—dropping sharply if blood is left at room temperature for even five hours before fixation—unlike other immune cell populations, which remain stable. For these reasons, unless specifically requested by a customer, we use SIGLEC-8 as a ‘dump gate’ to exclude eosinophils and increase confidence in the quantification of other granulocyte populations.
What about treating samples with heparin to improve eosinophil detection? It’s true—heparin can help, and we’ve seen improvements. But it comes with trade-offs. Heparin can negatively impact other functional markers like Ki67, TCF1, FOXP3, and Tbet.
If eosinophil quantification is important for your study, our team can discuss potential options. However, we do not report on eosinophil state markers, as their intracellular granules interfere with antibody binding.
We perform cytometry and specialize in peripheral blood mononuclear cells and fixed whole blood from humans.
While we don’t routinely process other sample types—such as dissociated tumors, bone marrow, or immune cells from humanized mice—we’re happy to discuss options.
We recommend collecting whole blood into standard vacutainers containing—in order of preference—heparin, citrate, or EDTA, and using our three-step TokuKit for on-site processing in clinical trials. This approach is faster, easier, and more cost-effective than PBMC isolation, with no centrifugation steps, and yields high-quality cytometry data with sample stability for months at -80°C.
We also work with viably cryopreserved PBMCs and whole blood samples that have undergone red blood cell lysis and fixation using PROT1 or Stable-Lyse, Stable-Store (from SmartTube).
We cannot accept whole blood collected in RNA or DNA isolation tubes, or serum samples, as these lack intact cells required for cytometry-based analysis.
We've used unsupervised clustering to analyze a 29M cell cytometry dataset to identify immune cell populations and subsets that correlate with response outcomes in melanoma patients treated with anti-PD1 therapy. In partnership with MGH, we identified a cluster of CD161+ CD4+ T cells that were assocaited with response and had a statistically significant increase in TBET and decrease in CCR7.
Paired with advanced quality control methods, we used unsupervised clustering to identify new immune cell populations and subsets who's frequencies and expression levels of functional proteins are relevant in response to treatment.
Check out our poster in our Articles page for more details!
We conducted an inter-kit precision study and observed an average coefficient of variation (CV) of 5.76% across all 33 immune cell populations analyzed. All populations were within the industry-standard 25% CV cutoff.
To assess inter-kit precision, we used samples from three donors, split one sample from each donor into three replicates, and fixed each of the nine samples using a separate TokuKit. We then analyzed the samples using our CLIA-validated, off-the-shelf, 40+ marker whole blood mass cytometry panel.
TokuKits typically have a minimum shelf life of 6 months when shipped.
The expiration date depends on the manufacturing date of the Stable-Lyse or PROT1 buffer included in the kit. Expiration dates are printed directly on each TokuKit and usually range from 6 months to 2 years.
Manual gating is a step-by-step process used in cytometry to identify immune cell populations and subsets based on the presence or absence of specific markers, often guided by published literature. For example, T cells express CD3 and B cells express CD19, so we can plot these two markers to identify each population (see Fig. A).
From there, we drill down into each population to define subsets. Within the CD3⁺ T cell gate, for instance, we can use CD4 and CD8 to distinguish CD4⁺ helper and CD8⁺ cytotoxic T cells. This process continues—gating further into CD4 or CD8 populations—until we’ve identified all relevant T cell subsets (see Fig. B). The same approach is used for B cells, NK cells, and other immune populations.
Our validation data show the minimum input for accurate detection of all major immune cell populations is 50K viable cells or ~700uL of fixed whole blood.
While these are the minimum volumes for detection of major immune cell population, we recommend starting with 1 million viable PBMCs or 2mL of whole blood for most samples.
It depends on the level of confidence you need and the amount of variability you can accept. Variability is measured by the coefficient of variation (CV), which is simply the standard deviation divided by the mean. A higher CV means more variability; a lower CV means more consistency.
Intuitively, common populations (e.g., 10%) require fewer total events to quantify accurately than rare ones (e.g., 0.001%).
This question has been addressed by Keeney et al., who proposed a simple formula to estimate the number of events needed for a given CV:
r = (100 / CV)²
where r is the number of events needed, and CV is the desired coefficient of variation.
For example, to measure a 0.1% population:
To get 10,000 events of a 0.1% population, you’d need 10 million total events. For 400 events, you'd need 400,000.
Intermediate monocytes (inMono) make up 0.47% of non-granulocytes in our internal dataset. To achieve a 5% CV, you’d need at least 188 inMono events. We collected 302—enough to achieve a 4.63% intra-run CV.
All immune populations measured using Teiko’s standard panel meet the accepted CV criteria of 25–30%.
For details and full calculations, read our article.
Functional markers are specific proteins that indicate the functional state or condition of a cell at a point in time. Some examples of different states of cells include activation, exhaustion, and proliferation. Some state markers on our off-the-shelf peripheral blood mononuclear cell (PBMC) panel are Ki67, a protein cells produce when actively replications/proliferating, and PD-1, a checkpoint protein expressed on activated and exhausted T cells.
For each cell analyzed, we measure the level of expression of all markers on our panel, including all functional markers. Data for functional markers included in our panels and our Dashboard are reported as frequency and a median channel value (MCV, for mass cytometry) or median fluorescence intensity (MFI, for spectral flow cytometry):
The frequency represents the proportion of cells within each cell subset expressing that functional marker above a set threshold. We say these cells are “positive” for this functional marker. This can answer the question “What proportion of T cells are exhausted?” with “37% of T cells are positive for PD-1 and therefore are likely exhausted.”
The MCV or MFI represents the intensity of the signal detected for a specific metal isotope or fluorochrome associated with a particular antibody on a cell. This can answer the question “Does the amount of PD-1 expressed on T cells increase or decrease with treatment?” with “The amount of PD-1 expressed on T cells decreases with treatment because the MCV value for PD-1 on T cells drops from 5.6 to 4.8 after treatment.” This type of marker expression level information is important since higher PD-1 protein expression can correlate with a more exhausted phenotype (PNAS 2013).
We evaluate functional marker expression as an MCV or MFI for a given marker in the (1) marker-positive cell population and (2) parent population. This allows you to quantify the level of expression within cells that are known to express the functional marker of interest and across the entire parent population, including cells with no or low expression of the functional marker.
We tailor, or adjust, gates between patients because each individual has a unique immune setpoint, meaning their immune cells may express slightly higher or lower levels of various protein markers at baseline.
This causes the populations to shift on the plots as you cycle between patients and the gates should be moved to accommodate this shift per-patient/per-subject. Once the gates are set for each patient, we maintain these gates between their time points or conditions. This ensures we are able to capture shifts in expression of functional markers across time and conditions based on each patient’s unique baseline.
In earlier versions of Teiko’s Dashboard, the “top gate” referred to the cell population used as the denominator to calculate the relative frequency of an immune cell subset.
Note that for current and future projects, there are no restrictions—you can display, calculate, and analyze any population or subset using any ancestor in the cell lineage.
However, historically, by default:
For example:
We use different top-level gates because if everything were measured as a percent of live cells, some populations would be too rare to visualize clearly. For instance, plasmablasts are about 0.22% of non-granulocytes, but only 0.09% of total leukocytes.
You can check a population’s top gate in two places:
We recommend within 5 hours of the blood draw into a sodium heparin, EDTA, or citrate blood collection tube.
The collected blood can remain at room temperature in the vacutainer for up to 5 hours before processing with the TokuKit. We found that 100% of immune cell subsets are preserved when TokuKits are processed within 5 hours, but that delay to 7 hours results in loss of several important immune lineages.
Generally speaking, the sooner the blood is processed with the TokuKit after blood draw the better. If possible, we recommend processing within 2-hours of blood collection when possible. You can read more about these findings here.
We first determined the optimal channel for each marker based on antigen abundance. Then, we tested five antibody concentrations in a serial dilution to identify the optimal staining concentration for each antibody.
Depending on the marker, healthy cells are either left unstimulated or stimulated with an appropriate biological agent, such as PHA or PMA. In some cases, cultured cells or patient samples are used to verify staining.
Spillover signal is assessed by examining signal in nearby and relevant channels. If spillover is detected, an antibody is moved to a different channel and/or staining concentration is adjusted.
To confirm the panel can detect all major immune cell populations, we measure the frequencies of key populations and subsets in PBMCs from at least three healthy donors.
We’ve tested and verified over 130 unique antibodies, and counting.
The full list can be found on our Custom Panel Development page of our website.
Our 33- and 41-marker custom panel backbones were originally inspired by work from Hartmann et al. We’ve made significant improvements since then.
We optimized antibody channel placement to allow easier customization, reduce spillover, and improve staining and gating of immune cell populations and functional subsets. We also performed intra- and inter-run precision testing on the 41-marker backbone, showing single-digit CVs for reproducibility.
Testing is still ongoing, as we are waiting for time to elapse. Here are the numbers to date:
Up to 15 months stable for mass cytometry.
R > 0.99: correlation in population frequencies between 1 week and 15 months
R = 0.95: correlation in functional marker subset frequencies
R = 0.93: correlation in protein expression level
And up to 1 month by spectral flow cytometry.
R = 1: correlation in population frequencies between 1 week and 15 months
R = 0.99: correlation in functional marker subset frequencies
R = 0.91: correlation in protein expression level
We show that the frequencies of immune cell populations quantified in whole blood and PBMC with are cytometry assays are comparable and highly correlated. Check out our webinar for more details: Whole blood or PBMCs for immune profiling?
Not for our spectral flow cytometry assays—but we do fix PBMCs before staining in our mass cytometry assays and see no impact on the dynamic range of most functional markers.
For mass cytometry, we fix PBMCs with paraformaldehyde (PFA) prior to staining, since our panels are optimized for fixed samples. Hartmann et al. (Cell Reports, 2019) compared PFA fixation before staining (PFA-stained) vs. after staining (live-stained) and found immune cell population frequencies to be highly similar, with an r = 0.94 correlation.
They also reported that PFA fixation doesn’t affect the dynamic range of most functional markers, though a few antigens—like CCR7 and CD11b—showed reduced staining. Overall, the authors concluded that PFA fixation before antibody staining has minimal impact on immune profiling.
2 weeks, typically, upon receipt of reagents including the antibody, metal, and samples. In some cases—such as when a special stimulation or cell culture step is needed for marker expression—testing and verification can take up to 4 weeks.
We communicate this in advance so you're aware of the expected turnaround time. No surprises!
Gating your samples is one of the most crucial parts of data analysis.
This example shows how a bad gating strategy can you lead down the wrong road, in this case thinking that you have 30% CD8+ T cells, when it reality this signal is contamination from stick cells, likely platelets and erythrocytes.
Gating might be straightforward for smaller, 3-8 color panels, but becomes highly complex for 20+ marker panels identifying over 30 immune populations.
Our gating starts by excluding debris, dead cells, platelets and red blood cells. In a next step, we remove Basophils, Eosinophils and other Granulocytes. This allows us to report the remaining immune cell frequencies as a percentage of non-Granulocytes, making it easy for you to compare percentage numbers between samples.
Check out the flow plots for an example of how our gating strategy helps identify clean populations of true CD4+ and CD8+ T-cells within CD3+ non-Granulocytes.
The TokuKit contains a blood collection vacutainer (typically Heparin or EDTA, depending on your study), and two small bottles with cell preservation reagent. Two disposable pipettes are included to transfer blood and samples from one bottle to the next.
The process is simple and completed within 20 minutes: blood is collected in the 4 mL vacutainer, 2 mL of blood is transferred to the bottle labeled “1,” inverted 10 times, and incubated at room temperature for 15 minutes—no machine shaking, no centrifuge. The blood mixture is then transferred to the bottle labeled “2,” inverted 10 times, and is ready to be stored at -80°C or shipped to a central lab or Teiko on dry ice.
Super simple. Way easier—and at least 10x fewer steps—compared to isolating PBMCs.
Each antibody on Teiko’s cytometry panels goes through a multi-step verification process to ensure that the selected concentration provides optimal marker detection without spilling into neighboring channels.
The process for any marker starts with a six-point concentration series, beginning at 6 µg/mL and performing two-fold dilutions down to 0.1875 µg/mL to create six distinct concentrations. For each concentration, we measure the median channel values (similar to median fluorescence intensity in flow cytometry) and the standard deviation of the channel values for each marker in the cell populations that express (positive population) and do not express (negative population) the given marker.
These measurements allow us to calculate a Stain Index (SI) for each concentration. The SI is a key metric in cytometry that measures an antibody’s ability to distinguish between positive and negative populations.
When comparing stain indices, the goal is typically to select the concentration with the highest value. However, when SIs are similar, we also assess separation visually using the dot plot in the gating scheme, spillover into the +1, -1, and +16 channels, and the median channel value of the positive population at each concentration.
Note: the +1 channel refers to the isotope with a molecular weight one unit higher than the metal isotope bound to the antibody; the -1 channel refers to one unit lower. We check the +16 channel to ensure the metal has not oxidized—this can happen. If it has, we re-conjugate the antibody to fresh metal.
Check out our Articles page for a webinar and examples of how we determine the optimal staining concentrations of Tbet (intracellular marker) on our panels.
Yes, we offer long-term sample storage at -80°C or in liquid nitrogen (LN2) for a subscription-based fee. Reach out to discuss options.
Each antibody on Teiko’s cytometry panels goes through a multi-step verification process to ensure that the selected concentration provides optimal marker detection without spilling into neighboring channels.
The process for any marker starts with a six-point concentration series, beginning at 6 µg/mL and performing two-fold dilutions down to 0.1875 µg/mL to create six distinct concentrations. For each concentration, we measure the median channel values (similar to median fluorescence intensity in flow cytometry) and the standard deviation of the channel values for each marker in the cell populations that express (positive population) and do not express (negative population) the given marker.
These measurements allow us to calculate a Stain Index (SI) for each concentration. The SI is a key metric in cytometry that measures an antibody’s ability to distinguish between positive and negative populations.
When comparing stain indices, the goal is typically to select the concentration with the highest value. However, when SIs are similar, we also assess separation visually using the dot plot in the gating scheme, spillover into the +1, -1, and +16 channels, and the median channel value of the positive population at each concentration.
Note: the +1 channel refers to the isotope with a molecular weight one unit higher than the metal isotope bound to the antibody; the -1 channel refers to one unit lower. We check the +16 channel to ensure the metal has not oxidized—this can happen. If it has, we re-conjugate the antibody to fresh metal.
Check out our Articles page for a webinar and examples of how we determine the optimal staining concentrations of CD25 (surface marker).
We completed rigorous testing as part of our CLIA registration process. Using PBMCs and whole blood samples from multiple healthy donors, we assessed intra-run, inter-run, inter-operator, inter-instrument, and inter-kit variance.
Full CLIA validation reports for our standard PBMC and whole blood assays—both mass cytometry and spectral flow—are available for download at teiko.bio.
Each antibody on Teiko’s cytometry panels goes through a multi-step verification process to ensure that the selected concentration provides optimal marker detection without spilling into neighboring channels.
The process for any marker starts with a six-point concentration series, beginning at 6 µg/mL and performing two-fold dilutions down to 0.1875 µg/mL to create six distinct concentrations. For each concentration, we measure the median channel values (similar to median fluorescence intensity in flow cytometry) and the standard deviation of the channel values for each marker in the cell populations that express (positive population) and do not express (negative population) the given marker.
These measurements allow us to calculate a Stain Index (SI) for each concentration. The SI is a key metric in cytometry that measures an antibody’s ability to distinguish between positive and negative populations.
When comparing stain indices, the goal is typically to select the concentration with the highest value. However, when SIs are similar, we also assess separation visually using the dot plot in the gating scheme, spillover into the +1, -1, and +16 channels, and the median channel value of the positive population at each concentration.
Note: the +1 channel refers to the isotope with a molecular weight one unit higher than the metal isotope bound to the antibody; the -1 channel refers to one unit lower. We check the +16 channel to ensure the metal has not oxidized—this can happen. If it has, we re-conjugate the antibody to fresh metal.
Check out our Articles page for a webinar and examples of how we determine the optimal staining concentrations of Granzyme B (cytoplasmic marker).
Blood samples processed with the TokuKit are stored long-term at -80°C. For short-term storage and transport, samples can be kept on dry ice. Samples can remain in the final Buffer 2 tube at -80°C until ready for cytometry analysis.
TokuKits can be ordered online at teiko.bio/tokukit. If you're ordering on behalf of a clinical trial site, you’ll have the option to bill the drug sponsor at checkout.
Currently, we do not support the addition of peptide-MHC tetramers into our panels.
We offer two versions of the TokuKit—one with a sodium heparin tube and one with an EDTA tube.
2 mL is ideal, as the buffer ratios are optimized to ensure proper fixation at that volume. However, there is some flexibility—the kits can support fixation of 2 mL ± 0.5 mL.
Yes. However, monoclonal antibodies are generally preferred, as they provide the most reproducible results with minimal lot-to-lot variability.
If a monoclonal antibody isn’t available or doesn’t work for a specific target, we can use a polyclonal antibody. In those cases, we restrict use to a single lot and order enough to support the entire study to avoid variability.
Yes, if samples arrive at our facility within 48 hours of the blood draw.
Blood stored at room temperature for 24 to 48 hours shows modest changes in marker expression, but dramatic changes by 72 hours.
Temperature fluctuations during shipping and delivery delays are other factors that can impact fresh blood samples. For this reason, we highly recommend fixing blood on-site using our TokuKit and shipping samples on dry ice to preserve marker expression and sample quality.
Yes, conditionally, and no.
Yes – transcription factors and other highly-expressed intracellular markers can easily be detected. Our backbone panels typically include transcription factors such as Foxp3, T-bet, and TCF1 and intracellular functional proteins like Granzyme B and Ki-67. We have also validated a number of other transcription factors like HIF1a, intracellular proteins like Perforin, and even metabolic markers like mTOR and GAPDH.
Conditionally – some cytokines, such as IFNg and TNFa, can be detected in activated cells without additional stimulation. However, most cytokines typically require cell stimulation in the presence of a secretion blocker (like Brefeldin A) to build up sufficient protein for detection. Our scientists can help you determine whether prior stimulation may be necessary depending on your cytokine of interest.
No – chemokines are often produced in low quantities and are not well-suited for cytometry-based detection. While chemokine production cannot be easily detected, chemokine receptor expression is detectable and can provide information about which immune subsets are responsive to chemokine gradients.
Teiko currently does not offer TokuKits (or other collection kits) for tissues other than blood. TokuKits may be adaptable for bone marrow or dissociated, homogenized tumor tissue, but this would require further testing.
TokuKits are available for purchase at teiko.bio/tokukits. Our team is available to discuss your study and provide feedback on the approach.
All antibodies are tested and verified in-house by our team. When commercially available, we use pre-conjugated antibodies. If conjugation is needed, our team performs it to ensure high performance and fast turnaround. All new antibodies—whether pre-conjugated or conjugated in-house—undergo in-house testing and verification before including in a panel.
Yes. The procedure is well-established on mass cytometry (https://pmc.ncbi.nlm.nih.gov/articles/PMC9991871/#S1) for PBMCs, live whole blood and fixed whole blood (after stimulation). We have the expertise to implement this sort of protocol.
We have additional experience in developing cytometry panels to detect levels of phosphorylated proteins that are outside of a signaling cascade. We’ve tested, developed and stained these in custom cytometry panels, including AML. If your team is interested, here’s a detailed overview: https://teiko.bio/antibody-verification/.
Yes, we can work with any purified antibody that is stored in a protein-free format (i.e. without BSA or other added proteins).
For surface markers, we can work with IgG or IgM antibodies.
For intracellular markers, we recommend using IgG antibodies as they tend to better penetrate into the cell. We do not work with Fab fragments due to incompatibility with our conjugation procedure.
Metal conjugation is performed in-house and the metal-conjugated antibody is tested and optimized on in-house PBMC samples. If your marker of interest is not expressed on PBMCs, a cell line or other sample type should be provided or can be purchased for panel validation.
Teiko currently does not offer TokuKits for non-human samples, although they are likely compatible.
TokuKits are available for purchase at teiko.bio/tokukits. Our team is available to discuss your study and provide feedback on the development of the TokuKit for non-human samples.
Yes, you get access to .FCS and Gating-ML files for your project.
FCS is the standard file format for flow cytometry data. Paired with GatingML, it allows you to review and reproduce the gating strategy applied to your project.
It’s likely possible, though we haven’t formally tested TokuKits for conventional (non-spectral) flow cytometry.
Two peer-reviewed publications (e.g., Nguyen et al.) have demonstrated that blood fixed using the Stable-Lyse/Stable-Store buffers used in our TokuKits can be stained and analyzed by conventional flow cytometry. These studies found that some markers are unaffected by fixation, while others are—highlighting the need for careful marker selection and validation when designing a panel.
Yes, please let us know when planning your project how much of each sample you would like us to return and where to ship the remaining samples. Thawed PBMC samples can be separated into aliquots and viably cryopreserved for return shipment.
No, samples processed by CyTOF cannot be recovered after acquisition, as the process destroys the cells.
Unclear—we haven’t tested this, and we’re not aware of any publications using TokuKit-fixed whole blood samples for scRNA-seq. If you're interested in this application, you would need to test compatibility independently.
TokuKits are available for purchase at teiko.bio/tokukits. Our team is available to discuss your study and provide feedback on the development of the TokuKit for scRNA-seq.
Yes.
We ask that customers review, make changes, and approve the gating scheme within 24 hours of receiving the sample and gating QC report. If gating is approved and later modified after analysis—requiring re-gating and re-analysis—an additional fee will apply.
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