Recent studies have shown that insulin autoantibodies occur in patients with newly diagnosed insulin-dependent diabetes mellitus (IDDM) before exogenous insulin treatment. Our study was designed to test the hypothesis that insulin autoantibodies, like cytoplasmic islet cell antibodies (ICAs), can identify individuals with ongoing autoimmune beta-cell destruction and increased risk of IDDM development. Insulin autoantibodies detected by use of a radioligand-binding assay were found in 1.4% of normal controls, 4% of first-degree relatives of IDDM patients, and in 37% of newly diagnosed IDDM patients. A strong positive correlation between insulin autoantibodies and ICAs was observed. HLA typing of insulin-autoantibody-positive first-degree relatives of IDDM patients, as well as in the general population, revealed a strong association with HLA-DR3 and/or-DR4, suggesting that insulin autoantibodies are restricted to persons genetically susceptible to IDDM. In an ongoing study of beta-cell function in ICA-positive nondiabetic individuals, the additional presence of insulin autoantibodies significantly increased the likelihood of beta-cell dysfunction. After intravenous glucose stimulation, insulinopenia was present in 70% of ICA and insulin-autoantibody-positive individuals in contrast to only 23% of ICA-positive, insulin-autoantibody-negative persons. These data document a significant association between insulin autoantibodies and ICAs and support the contention that insulin autoantibodies, like ICAs, are markers of ongoing beta-cell destruction.

Background Aims Multi-center cellular therapy clinical trials require the establishment and implementation of standardized cell processing protocols and associated quality control mechanisms. The aims here were to develop such an infrastructure in support of the Cardiovascular Cell Therapy Research Network (CCTRN) and to report on the results of processing for the first 60 patients. Methods Standardized cell preparations, consisting of autologous bone marrow mononuclear cells, prepared using the Sepax device were manufactured at each of the five processing facilities that supported the clinical treatment centers. Processing staff underwent centralized training that included proficiency evaluation. Quality was subsequently monitored by a central quality control program that included product evaluation by the CCTRN biorepositories. Results Data from the first 60 procedures demonstrate that uniform products, that met all release criteria, could be manufactured at all five sites within 7 hours of receipt of the bone marrow. Uniformity was facilitated by use of the automated systems (the Sepax for processing and the Endosafe device for endotoxin testing), standardized procedures and centralized quality control. Conclusions Complex multicenter cell therapy and regenerative medicine protocols can, where necessary, successfully utilize local processing facilities once an effective infrastructure is in place to provide training, and quality control.

Conventional flow cytometric methods for CD34+ cell counting may be affected by the high number of nucleated red blood cells or nonviable cells in cord blood and its products. We developed a simple flow cytometric no-wash procedure that avoids these shortcomings because it provides absolute CD34+ cell counts and assesses cell viability. Samples were incubated with phycoerythrin (PE)-labeled anti-CD34 (Becton Dickinson Immunocytometry Systems [BD], San Jose, CA) and peridinin chlorophyll protein (PerCP)-labeled anti-CD45 (BD) in bead-containing TRUCOUNT tubes (BD). After red cell lysis with a fixative-free reagent, the impermeant nucleic acid dye YO-PRO-1 (Molecular Probes, Eugene, OR) was added and samples were analyzed on a single-laser FACSCalibur (BD). A comparison with the ProCOUNT progenitor cell assay (BD) in 57 samples revealed excellent correlation of results (r = 0.98, intercept -0.2 cells/microl, slope 1.01). Precision studies conveyed coefficients of variation of 6.4 and 8.9% at concentrations of 35 and 16 CD34+ cells/microl, respectively. In untreated and leukocyte-enriched cord blood 4.5+/-3.8% of CD34+ cells were stained by YO-PRO-1, representing apoptotic or necrotic cells. In post-thawing cryopreserved samples this number increased to 10.4+/-5.5%. Isotype controls showed very low blank values of viable cells (0.1+/-0.4 cells/microl, maximum 2.4) and seemed unnecessary. We found no washing-related alteration of results in 35 samples, indicating that the method may also be performed with cell washing. Replacing YO-PRO-1 with TO-PRO-3 facilitated four-color analysis of subpopulations of viable CD34+ cells on a FACSCalibur equipped with a second (diode) laser. We found the proposed method to be a rapid, efficient, and flexible procedure that improved validity of CD34+ cell counts.