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Background Aims: Wide acceptance of the colony-forming unit (CFU) assay as a reliable potency test for stem cell products is hindered by poor inter-laboratory reproducibility. The goal of this study was to ascertain current laboratory practices for performing the CFU assay with an eye towards identifying practices that could be standardized to improve overall reproducibility., Methods: A survey to evaluate current laboratory practices for performing CFU assays was designed and internationally distributed., Results: There were 105 respondents to the survey, of whom 68% performed CFU assays. Most survey recipients specified that an automated rather than a manual cell count was performed on pre-diluted aliquots of stem cell products. Viability testing methods employed various stains, and when multiple sites used the same viability stain, the methods differed. Cell phenotype used to prepare working cell suspensions for inoculating the CFU assay differed among sites. Most respondents scored CFU assays at 14-16 days of incubation, but culture plates were read with various microscopes. Of 57 respondents, 42% had not performed a validation study or established assay linearity. Only 63% of laboratories had criteria for determining if a plate was overgrown with colonies., Conclusions: Survey results revealed inconsistent inter-laboratory practices for performing the CFU assay. The relatively low number of centers with validated CFU assays raises concerns about assay accuracy and emphasizes a need to establish central standards. The survey results shed light on numerous steps of the methodology that could be targeted for standardization across laboratories., (Copyright © 2013 International Society for Cellular Therapy. All rights reserved.)

Background: The stroma-based long-term culture is the assay of choice when a functional detection of primitive hematopoietic cells in vitro is sought. However, different stromal cell lines varying in supporting capacity have been raised and applied in different laboratories, resulting in a wide range in published frequencies of LTCIC alternative CAFC., Methods: In order to identify the most suitable stromal source in terms of supportive capacity, reproducibility, and ease of handling, we have compared some of the most commonly employed murine cell lines to human bone marrow stroma in secondary long-term culture set-ups., Results: Seeking an approximation to the supportive capacity of human BM stroma we found the FBMD-1 cell line supplemented with G-CSF and IL-3 superior to FBMD-1 cells alone, and to M2-10B4 and Sl/Sl cells. Moreover, in co-cultures of CD34(+) cells and the FBMD-1 line, we found week 5 CAFC content highly reproducible (50.5 +/- 6.66 - 54.6 +/- 7.07/10(4) plated cells, p value > 0.95) and the assay was suitable for inter-individual comparison in a clinical setting. In fact, the week 5 CAFC results were even more reproducible than those of the CFU assays (CV 0.03 for the CAFC assay versus 0.13-0.33 for the CFU assays). On the other hand, when extending the culture period to 8 weeks, the cobblestone area formation was best maintained by human BM stroma and the high reproducibility in CAFC enumeration in cultures supported by the FBMD-1 was lost., Discussion: Among the stromal cell sources tested, the FBMD-1 line was found to be superior in terms of ease of handling and week 5 CAFC reproducibility. However, this robustness could not be extended to week 8 CAFC.

We report here the development of a rapid and quantitative method for measuring in vitro T cell transformation by human T cell leukemia viruses type I (HTLV-I) and type II (HTLV-II). This method is based on our finding that cocultivation of lethally irradiated HTLV-producing cells with peripheral blood lymphocytes (PBLs) preactivated for 24 hours with phytohemagglutinin and interleukin-2 (IL-2) induces colony formation in methylcellulose-containing medium. Colonies of about 200 cells can be clearly distinguished from background aggregates within four to six days after cocultivation. These colonies gradually increase in size and reach 300 to 1,000 cells within 14 days after cocultivation. Cells of these colonies were infected, as evidenced by expression of viral p19 antigen and the presence of HTLV proviral sequences. These cells proved to be transformed in terms of IL-2-independent continuous growth in liquid medium. Colony formation was found to depend in a linear fashion upon the percentage of the infected cells present in the irradiated cell population and is sufficiently sensitive to detect as few as 1% of virus-producing cells.

In mutation and cell transformation assays, it has long been recognized that the common practice of using different numbers of cells on dishes with or without selective conditions creates a source of bias in mutant fraction determination. This is simply because colony formation may be enhanced or suppressed at higher initial cell densities, depending on the assay and agent tested. We propose a solution that consists of the inclusion of an experimentally distinguishable population of cells as an internal standard for colony-forming ability at the high cell density required for detection of rare variants. This method is found to be highly satisfactory for use in measuring mutation to 6-thioguanine resistance in a diploid human B lymphoblast line. For treatment with anti-2,3-dihydroxy-1,10b-epoxy-1,2,3-trihydrofluoranthene (FDE), N-methyl-N'-nitro-N-nitrosoguanidine (MNNG), and 4-nitroquinoline-oxide (4NQO), the calculated induced mutant fractions using the internal-standard method were significantly lower than those calculated using the conventional low-density-plating efficiency method. The results of these experiments and our analysis lead us to conclude that this approach is applicable to all single cell mutation or transformation assays and is a necessary feature of assays in which an accurate knowledge of the fraction of rare variants is required.