Human Gene Therapy Methods最新文献

Recombinant adeno-associated virus (rAAV) vectors have recently been widely utilized for in in vivo gene therapy. The clinical dose definition of AAV vector requires the exact quantification as starting doses and for dose-escalation studies. Vector genome (vg) copies measured by quantitative PCR (qPCR) are commonly used for rAAV vector titration, and rAAV vector plasmids DNA is often used for qPCR standards, although the rAAV reference standard materials (RSMs) for serotypes 2 and 8 (rAAV2RSM and rAAV8RSM) are available from American Type Culture Collection. However, qPCR-based determination of the AAV vg is affected by the selection of the qPCR standard and the amplification target sites. In this study, we have developed a new PCR method, two-dimensional droplet digital PCR (2D ddPCR), for the absolute quantitation of target DNA and for evaluating the stability of the rAAV vector. The number of vg copies of rAAV2RSM determined by qPCR dramatically changed when standard plasmid DNAs with different conformations were treated with restriction enzymes, suggesting that qPCR amplification is significantly affected by the secondary structure of the standard. In contrast, the number of vg copies determined by ddPCR was unaffected by using primer probes for different positions of target sites or by the secondary structure conformation of the vg. Furthermore, the integrity of the AAV vg can be monitored using 2D ddPCR with fluorescein- and hexachloro-6-carboxy-fluorescine-labeled probes targeting different positions in the same rAAV genome. The titer of intact rAAV was highly correlated with rAAV activity in an accelerated (37°C) stability study. 2D ddPCR is a useful tool for rAAV vector quantitation and quality evaluation.

The study of human cytomegalovirus (HCMV) has for long been challenging due to the inability of clinical strains to efficiently proliferate in vitro until adaptive mutations occur. These mutations lead to strains that differ considerably from clinical isolates, many of them showing altered cell tropism, a decrease in cell association and higher susceptibility to an innate immune response. These problems were recently solved by the use of bacterial artificial chromosome (BAC) vectors that allow for the conservation of an intact HCMV genome. Other characteristics that render HCMV difficult for in vitro study are related to its slow replication rate that leads to some constraints in its titration. During the cloning of HCMV into BAC vectors, many groups additionally inserted a fluorescent tag to facilitate the virus characterization. However, the methods used for titration of HCMV-BAC stocks are still relaying on the standard methods that are expensive and/or time consuming. In this study, we assessed the possibility of viral titration by fluorescence-activated cell sorting (FACS), making use of the fluorescent tags that many of the HCMV-BACs hold. We compared viral titers obtained by immunohistochemistry with FACS, a faster and inexpensive technique. We showed that viral titers are comparable using the techniques already mentioned, and that titration by FACS is an efficient, fast, and cost-effective method. The establishment of viral titration of BAC vectors by FACS can further simplify the study of HCMV.

Scalable lentiviral vector (LV) manufacturing is vital for successful commercialization of LV-based gene and cell therapy products. Accordingly, efforts are currently focused on developing and adapting technologies to address both upstream and downstream production bottlenecks. To overcome the limitations of current upstream processes, researchers are now favoring the use of bioreactors over traditional two-dimensional culture platforms. Bioreactors provide many advantages for manufacturing biomolecules, including process automation, tight regulation of production conditions, reduced labor input, and higher productivity potential. This study describes a transient LV production strategy employing a single-use, packed-bed bioreactor vessel. Functional LV titers in the 10⁶ TU/mL range were achieved, and after concentration yields of up to 10⁹ TU/mL were attained. This proof of principle study demonstrates that LV can be successfully produced in a packed-bed system. With further optimization, a packed-bed bioreactor could offer a potential scale-out solution for LV manufacturing.

Innate immune signals that promote B cell responses in gene transfer are generally ill-defined. In this study, we evaluate the effect of activating endosomal Toll-like receptors 7, 8, and 9 (TLR7, TLR7/8, and TLR9) on antibody formation during muscle-directed gene therapy with adeno-associated virus (AAV) vectors. We examined whether activation of endosomal TLRs, by adenine analog CL264 (TLR7 agonist), imidazolquinolone compound R848 (TLR7/8 agonist), or class B CpG oligodeoxynucleotides ODN1826 (TLR9 agonist), could augment antibody formation upon intramuscular administration of AAV1 expressing human clotting factor IX (AAV1-hFIX) in mice. The TLR9 agonist robustly enhanced antibody formation by the 1st week, thus initially eliminating systemic hFIX expression. By contrast, the TLR7 and TLR7/8 agonists did not markedly promote antibody formation, or significantly reduce circulating hFIX. We concurrently investigated the effects of these TLR agonists during muscle gene transfer on mature B cells and dendritic cells (DCs) in the draining lymph nodes including conventional DCs (CD11b⁺ or CD8α⁺ cDCs), monocyte-derived dendritic cells (moDCs), and plasmacytoid dendritic cells (pDCs). Only TLR9 stimulation caused a striking increase in the frequency of moDCs within 24 h. The TLR7/8 and TLR9 agonists activated pDCs, both subsets of cDCs, and mature B cells, whereas the TLR7 agonist had only mild effects on these cells. Thus, these TLR ligands have distinct effects on DCs and mature B cells, yet only the TLR9 agonist enhanced the humoral immune response against AAV-expressed hFIX. These new findings indicate a unique ability of certain TLR9 agonists to stimulate B cell responses in muscle gene transfer through enrichment of moDCs.

Duchenne muscular dystrophy (DMD) is a severe type of X-linked recessive degenerative muscle disease caused by mutations in the dystrophin (DMD) gene on the X chromosome. The DMD gene is complex, consisting of 79 exons, and mutations cause changes in the DMD mRNA so that the reading frame is altered, and the muscle-specific isoform of the dystrophin protein is either absent or truncated with variable residual function. The emerging CRISPR-Cas9-mediated genome editing technique is being developed as a potential therapeutic approach to treat DMD because it can permanently replace the mutated dystrophin gene with the normal gene. Prenatal DNA testing can inform whether the female fetus is a carrier of DMD, and the male fetus has inherited a mutation from his mother (50% chance of both). This article summarizes the present status of current and future treatments for DMD.

In cellular immunotherapies, natural killer (NK) cells often demonstrate potent antitumor effects in high-risk cancer patients. But Good Manufacturing Practice (GMP)-compliant manufacturing of clinical-grade NK cells in high numbers for patient treatment is still a challenge. Therefore, new protocols for isolation and expansion of NK cells are required. In order to attack resistant tumor entities, NK cell killing can be improved by genetic engineering using alpharetroviral vectors that encode for chimeric antigen receptors (CARs). The aim of this work was to demonstrate GMP-grade manufacturing of NK cells using the CliniMACS^® Prodigy device (Prodigy) with implemented applicable quality controls. Additionally, the study aimed to define the best time point to transduce expanding NK cells with alpharetroviral CAR vectors. Manufacturing and clinical-scale expansion of primary human NK cells were performed with the Prodigy starting with 8-15.0 × 10⁹ leukocytes (including 1.1-2.3 × 10⁹ NK cells) collected by small-scale lymphapheresis (n = 3). Positive fraction after immunoselection, in-process controls (IPCs), and end product were quantified by flow cytometric no-wash, single-platform assessment, and gating strategy using positive (CD56/CD16/CD45), negative (CD14/CD19/CD3), and dead cell (7-aminoactinomycine [7-AAD]) discriminators. The three runs on the fully integrated manufacturing platform included immunomagnetic separation (CD3 depletion/CD56 enrichment) followed by NK cell expansion over 14 days. This process led to high NK cell purities (median 99.1%) and adequate NK cell viabilities (median 86.9%) and achieved a median CD3+ cell depletion of log -3.6 after CD3 depletion and log -3.7 after immunomagnetic CD3 depletion and consecutive CD56 selection. Subsequent cultivation of separated NK cells in the CentriCult^® chamber of Prodigy resulted in approximately 4.2-8.5-fold NK cell expansion rates by adding of NK MACS^® basal medium containing NK MACS^® supplement, interleukin (IL)-2/IL-15 and initial IL-21. NK cells expanded for 14 days revealed higher expression of natural cytotoxicity receptors (NKp30, NKp44, NKp46, and NKG2D) and degranulation/apoptotic markers and stronger cytolytic properties against K562 compared to non-activated NK cells before automated cultivation. Moreover, expanded NK cells had robust growth and killing activities even after cryopreservation. As a crucial result, it was possible to determine the appropriate time period for optimal CAR transduction of cultivated NK cells between days 8 and 14, with the highest anti-CD123 CAR expression levels on day 14. The anti-CD123 CAR NK cells showed retargeted killing and degranulation properties against CD123-expressing KG1a target cells, while basal cytotoxicity of non-transduced NK cells was determined using the CD123-negative cell line K562. Time-lapse imaging to monitor redirected effector-to-target con