From the perspective of a pilot clinical gene therapy trial for Wiskott-Aldrich syndrome (WAS), we implemented a process to produce a lentiviral vector under good manufacturing practices (GMP). The process is based on the transient transfection of 293T cells in Cell Factory stacks, scaled up to harvest 50 liters of viral stock per batch, followed by purification of the vesicular stomatitis virus glycoprotein-pseudotyped particles through several membrane-based and chromatographic steps. The process leads to a 200-fold volume concentration and an approximately 3-log reduction in protein and DNA contaminants. An average yield of 13% of infectious particles was obtained in six full-scale preparations. The final product contained low levels of contaminants such as simian virus 40 large T antigen or E1A sequences originating from producer cells. Titers as high as 2 × 10(9) infectious particles per milliliter were obtained, generating up to 6 × 10(11) infectious particles per batch. The purified WAS vector was biologically active, efficiently expressing the genetic insert in WAS protein-deficient B cell lines and transducing CD34(+) cells. The vector introduced 0.3-1 vector copy per cell on average in CD34(+) cells when used at the concentration of 10(8) infectious particles per milliliter, which is comparable to preclinical preparations. There was no evidence of cellular toxicity. These results show the implementation of large-scale GMP production, purification, and control of advanced HIV-1-derived lentiviral technology. Results obtained with the WAS vector provide the initial manufacturing and quality control benchmarking that should be helpful to further development and clinical applications.

Lentiviral vectors are effective tools for gene transfer and integrate variable numbers of proviral DNA copies in variable proportions of cells. The levels of transduction of a cellular population may therefore depend upon experimental parameters affecting the frequency and/or the distribution of vector integration events in this population. Such analysis would require measuring vector copy numbers (VCN) in individual cells. To evaluate the transduction of hematopoietic progenitor cells at the single-cell level, we measured VCN in individual colony-forming cell (CFC) units, using an adapted quantitative PCR (Q-PCR) method. The feasibility, reproducibility and sensitivity of this approach were tested with characterized cell lines carrying known numbers of vector integration. The method was validated by correlating data in CFC with gene expression or with calculated values, and was found to slightly underestimate VCN. In spite of this, such Q-PCR on CFC was useful to compare transduction levels with different infection protocols and different vectors. Increasing the vector concentration and re-iterating the infection were two different strategies that improved transduction by increasing the frequency of transduced progenitor cells. Repeated infection also augmented the number of integrated copies and the magnitude of this effect seemed to depend on the vector preparation. Thus, the distribution of VCN in hematopoietic colonies may depend upon experimental conditions including features of vectors. This should be carefully evaluated in the context of ex vivo hematopoietic gene therapy studies.

Improved, human-based packaging cell lines allow the production of high-titer, RCR-free retroviral vectors. The utility of these cell lines for the production of clinical grade vectors critically depends on the definition of optimal conditions for scaled-up cultures. In this work, a clone derived from the TE Fly GALV packaging cell (Duisit et al. Hum. Gene Ther. 1999, 10, 189) that produces high titers of a lacZ containing retroviral vector with a Gibbon Ape Leukemia Virus envelope glycoprotein was used. This clone can produce (2-5) x 10(6) PFU cm(-3) in small scale cultures and has been evaluated for growth and vector production in different reactor systems. The performances of fixed bed reactors [CellCube (Costar) and Celligen (New Brunswick)] and stirred tank reactors [microcarriers and clump cultures] were compared. The cells showed a higher apparent growth rate in the fixed bed reactor systems than in the suspension systems, probably as a result of the fact that aggregation and/or formation of clumps led to a reduced viability and reduced growth of cells in the interior of the clumps. As a consequence, the final cell density and number were in average 3- to 7-fold higher in the fixed bed systems in comparison to the suspension culture systems. The average titers obtained ranged from 0.5 to 2.1 x 10(7) PFU cm(-3) for the fixed bed and microcarrier systems, while the clump cultures produced only (2-5) x 10(5) PFU cm(-3). The differences in titers reflect cell densities as well as specific viral vector production rates, with the immobilization and microcarrier systems exhibiting an at least 10-fold higher production rate in comparison to the clump cultures. A partial optimization of the culture conditions in the Celligen fixed bed reactor, consisting of a 9-fold reduction of the seeding cell density, led to a 5-fold increased vector production rate accompanied by an average titer of 3 x 10(7) PFU cm(-3) (maximum titer (4-5) x 10(7) PFU cm(-3)) in the fixed bed reactor. The performance evaluation results using mathematical models indicated that the fixed bed bioreactor has a higher potential for retroviral vector production because of both the higher reactor productivity and the lower sensitivity of productivity in relation to the changes in final retrovirus titer in the range of 3 x 10(6) to 15 x 10(6) PFU cm(-3).

Neutralizing antibodies directed against adeno-associated virus (AAV) are commonly found in humans. In seropositive subjects, vector administration is not feasible as antibodies neutralize AAV vectors even at low titers. Consequently, a relatively large proportion of humans is excluded from enrollment in clinical trials and, similarly, vector redosing is not feasible because of development of high-titer antibodies following AAV vector administration. Plasmapheresis has been proposed as strategy to remove anti-AAV antibodies from the bloodstream. Although safe and relatively effective, the technology has some limitations mainly related to the nonspecific removal of all circulating IgG. Here we developed an AAV-specific plasmapheresis column which was shown to efficiently and selectively deplete anti-AAV antibodies without depleting the total immunoglobulin pool from plasma. We showed the nearly complete removal of anti-AAV antibodies from high titer purified human IgG pools and plasma samples, decreasing titers to levels that allow AAV vector administration in mice. These results provide proof-of-concept of a method for the AAV-specific depletion of neutralizing antibodies in the setting of in vivo gene transfer.