Development of highly sensitive methods for defining off target mutations to enable safe gene editing of haematopoietic cells for transplantation

RNA引导内切酶（RGENs），如CRISPR-Cas9系统，已经彻底改变了我们编辑基因组的能力，因为它们允许我们按照RNA向导定义的序列切割和编辑基因组，RNA向导可以快速设计和制造。这对分子生物学产生了巨大的影响；允许编辑各种不同细胞类型和生物体的基因组。下一个挑战是利用这项技术编辑人类细胞的基因组来治疗疾病。造血是该技术将成功应用于临床的首批领域之一，因为造血干细胞（HSC）移植已经进行了40多年，收集、纯化和移植人类HSC的设施也已经建立起来。在患者中使用基因组编辑技术的主要障碍之一是在编辑过程中由意外突变引起恶性转化的潜在风险。为了将基因组编辑技术改进到可以安全地用于人体试验的程度，开发出用于确定经过基因组编辑的原代细胞突变的检测方法至关重要。这些脱靶效应需要以无偏倚的全基因组方式准确定义，并且检测需要高度敏感，以便能够检测到罕见的脱靶突变，因为恶性肿瘤源于单细胞的克隆转化。迄今为止，已经描述了几种确定脱靶效应的方法，但没有一种技术能够敏感而准确地定义原代细胞中的突变。这是这些技术临床应用的一个关键障碍，因为没有快速和经济有效的方法来比较各种不同的基因组编辑技术的准确性，将很难开发出用于人体试验的安全编辑方法。我将开发一种新方法，初步识别裸DNA中体外RGEN的所有潜在位点（这已被证明是对潜在脱靶活性位点的高度预测）。随后，这些位点将使用生物素化寡核苷酸进行深度测序，以在体外鉴定的目标位点捕获DNA。使用这种方法，它应该是相对简单的，使分析的灵敏度至少比现有的最佳方法高100倍。我还将使用这种方法寻找病毒载体和模板序列意外插入的罕见位点。在编辑过程中发生的突变的总体负担也将被调查。这将通过在编辑过程前后对流式细胞术纯化的造血干细胞和祖细胞进行全基因组测序来完成。方法将被用来减少测序错误，软件将被编写来定义从测序衍生的序列的可变性突变的总体负担。这些方法将用于优化用于地中海贫血和镰状细胞病治疗的两种基因组编辑模型，以用于潜在的临床应用。首先，在宿主实验室开发了一个模型，旨在通过删除α珠蛋白基因上的关键转录因子结合位点来治愈输血依赖性HbE -地中海贫血患者（约占输血依赖性地中海贫血的50%）。此外，将建立一种既定的方法，该方法使用RGEN插入DNA模板来原位校正β珠蛋白基因，同时表达细胞表面标记物，从而纯化校正后的细胞。这将允许我测试在使用rgen编辑细胞的两种主要策略上量化脱靶效应的方法。

RNA guided endonucleases (RGENs), such as the CRISPR-Cas9 system have revolutionised our ability to edit the genome because they allow us to cut and edit the genome at the sequence defined by a RNA guide, which can be rapidly designed and manufactured. This has had a huge impact on molecular biology; allowing the genome of a wide variety of different cell types and organisms to be edited. The next challenge is to use this technology to edit the genome of human cells to treat disease. Haematopoiesis is one of the first areas in which this technology will be successfully implemented in the clinic because haematopoietic stem cell (HSC) transplantation has been performed for over 40 years and facilities for collecting, purifying and transplanting human HSCs are well established. One of the major barriers to using genome editing technology in patients is the potential risk of malignant transformation from unintended mutations during the editing process. In order to improve genome editing techniques to the point at which they can be safely used in human trials it is critical that assays are developed to define mutations in primary cells that have undergone genome editing. These off target effects need to be defined accurately in an unbiased genome wide fashion and the assays need to be highly sensitive so that rare off target mutations can be detected because malignancy arises from clonal transformation of single cells. To date, several methods have been described for determining off target effects but none of the techniques is able to define mutations sensitively and accurately in primary cells. This is a key stumbling block to the clinical application of these techniques because without rapid and cost effective methods to compare the accuracy of the huge variety of different techniques available for genome editing it will be difficult to develop safe methods of editing for use in human trials. I will develop a novel method that initially identifies all potential sites of in vitro RGEN in naked DNA (this has previously been shown to be a highly predictive of potential sites of off target activity). Subsequently these sites will be sequenced at great depth using biotinylated oligonucleotides designed to capture DNA at the target sites identified in vitro. Using this it should be relatively straightforward to make the assay is at least 100 times more sensitive than the best available methods. I will also use this approach to look for rare sites of unintended insertion of viral vectors and template sequences. The overall burden of mutations that occur during the editing process will also be investigated. This will be done by performing whole genome sequencing on haematopoietic stem and progenitor cells purified by flow cytometry both before and after the editing process. Methods will be used to minimise sequencing errors and software will be written to define the overall burden of mutations from the variability of the sequences derived from sequencing. These methods will be used to optimise two models of genome editing for the treatment of thalassaemia and sickle cell disease for potential clinical use. First a model has been developed in the host laboratory which aims to cure patients with transfusion dependent HbE beta thalassaemia, which accounts for around 50% of transfusion dependent thalassaemia, by deleting key transcription factor binding sites at the alpha globin gene. In addition an established method will be set up, which uses an RGEN to insert a DNA template to correct the beta globin gene in situ and simultaneously express a cell surface marker that allows purification of corrected cells. This will allow me to test the methods for quantification of off target effects on the two main strategies for using RGENs for editing cells.