CRISPR/Cas9介导的低产尿素黄酒酵母工程菌的构建

通过代谢工程改造构建低产尿素的酿酒酵母工程菌,从根源上减少黄酒发酵液中尿素的含量及氨基甲酸乙酯(ethyl carbamate,EC)的形成。该研究利用融合PCR构建DUR3过表达组件"HOL-PGK1p-DUR3-PGK1tHOR",通过CRISPR/Cas9介导的基因组编辑技术转化酿酒酵母S. cerevisiae NaDUR1,2-Δcar1,在敲除CAR1和过表达DUR1,2基因的基础上过表达DUR3基因,获得工程菌S. cerevisiae NaDUR1,2/DUR3-Δcar1。实验室黄酒发酵实验结果表明,与亲本菌株S. cerevisiae Na相比,工程菌S. cerevisiae NaDUR1,2/DUR3-Δcar1所酿黄酒发酵液中尿素含量降低了92. 1%,EC含量降低了58. 6%;与出发菌株S. cerevisiae NaDUR1,2-Δcar1相比,工程菌S. cerevisiae NaDUR1,2/DUR3-Δcar1所酿黄酒发酵液中尿素含量降低了43. 4%,EC含量降低了16. 2%。过表达DUR3的工程菌S. cerevisiae NaDUR1,2/DUR3-Δcar1具有"尿素吸收"的能力,减少EC的形成。借助CRISPR/Cas9系统,构建的酵母工程菌无外源抗性基因的引入,具有工业化应用的潜在可能性。     

CRISPR/Cas9介导烟草多基因编辑体系的应用

CRISPR/Cas9是一种新型的基因组定向编辑技术,近年来在烟草基因组的定向编辑中得到了广泛应用。CRISPR/Cas9介导的多基因编辑技术在许多物种中表现出很大的潜力,为了探索CRISPR/Cas9介导烟草多基因编辑的技术体系,本文针对烟草PCS1、HMA2、eIF4E、TOM3、TOM1 5个不同性状相关的基因构建了多靶点敲除的CRISPR/Cas9系统,并借助农杆菌介导的遗传转化技术转化烟草品种K326。对阳性植株的筛选、靶位点基因组的PCR扩增与测序分析表明,构建的CRISPR/Cas9多基因编辑系统成功转化到烟草中,能够同时靶向突变5个基因。5个基因同时突变的检出率为53.8%,单基因的突变检出率在76.9%与92.3%之间。进一步的脱靶检测分析显示,在所预测脱靶的候选位点上均未发生脱靶现象。本文构建的CRISPR/Cas9多基因编辑系统可将多个基因进行有效突变,为烟草基因功能研究和基于CRISPR/Cas9技术的多性状改良奠定了基础。      

基于CRISPR/Cas9系统的金针菇基因组编辑载体构建

采用高效基因编辑系统CRISPR/Cas9,构建金针菇基因Mads-8敲除载体及转化体系。以转录组数据为依据,通过分析基因Mads-8序列信息,选择高效的靶位点序列,同时加入筛选标记基因hph构建过渡载体sgRNA-T,通过菌落PCR鉴定和测序来验证靶位点序列是否正确插入。以表达载体pg Fvs-cas9为框架,酶切连接后构建重组敲除载体pg Fvs-Cas9-gRNA,PCR和酶切鉴定敲除载体。再以PEG介导的金针菇原生质体转化表达载体,在潮霉素抗性平板上筛选拟转化子,以拟转化子基因组DNA为模板扩增Cas9基因。结果显示,靶位点序列成功插入敲除载体且序列正确,重组质粒成功转化进金针菇的基因组。对金针菇基因组敲除载体的构建,为后续金针菇相关基因的功能验证及育种研究提供载体材料及理论依据。      

基于CRISPR/Cas9系统对乳酸乳球菌进行绿色荧光蛋白标记

乳酸乳球菌是治疗剂在体内运送的良好载体,研究其在体内的真实运送情况需对其进行标记。该实验利用CRISPR/Cas9系统对乳酸乳球菌(Lactococcus lactis)NZ9000进行增强型绿色荧光蛋白(enhanced green fluorescent protein, eGFP)标记,用于研究乳酸乳球菌在体内的运送,评价其作为益生菌的功能。基于该实验室已构建的乳酸乳球菌CRISPR/Cas9编辑质粒pLL25构建重组质粒pYSH,其上携带eGFP及同源臂,电转入乳酸乳球菌NZ9000感受态细胞中,将基因组中的乳酸脱氢酶基因(ldh)替换为绿色荧光蛋白基因,从而使Lactococcus lactis NZ9000获得标记,表达绿色荧光蛋白。对绿色荧光标记的Lactococcus lactis NZ9000突变株,酶标仪定量分析eGFP的表达强度。荧光强度定量分析结果表明,在乳酸乳球菌不同生长阶段,eGFP基因均能稳定表达。     

基于CRISPR/Cas9系统对乳酸乳球菌进行绿色荧光蛋白标记

乳酸乳球菌是治疗剂在体内运送的良好载体,研究其在体内的真实运送情况需对其进行标记。该实验利用CRISPR/Cas9系统对乳酸乳球菌(Lactococcus lactis)NZ9000进行增强型绿色荧光蛋白(enhanced green fluorescent protein, eGFP)标记,用于研究乳酸乳球菌在体内的运送,评价其作为益生菌的功能。基于该实验室已构建的乳酸乳球菌CRISPR/Cas9编辑质粒pLL25构建重组质粒pYSH,其上携带eGFP及同源臂,电转入乳酸乳球菌NZ9000感受态细胞中,将基因组中的乳酸脱氢酶基因(ldh)替换为绿色荧光蛋白基因,从而使Lactococcus lactis NZ9000获得标记,表达绿色荧光蛋白。对绿色荧光标记的Lactococcus lactis NZ9000突变株,酶标仪定量分析eGFP的表达强度。荧光强度定量分析结果表明,在乳酸乳球菌不同生长阶段,eGFP基因均能稳定表达。     

CRISPR/Cas生物传感器检测食源性病原体的研究进展

食源性病原体引起的食源性疾病严重威胁着人类的健康,灵敏且快速检测食源性病原体成为预防疾病发生的重要手段。生物传感器具有灵敏度高,无需富集即可实时定量,易于现场检测等优点。CRISPR/Cas系统与生物传感器结合可提高生物传感器检测的灵敏度和准确性。本文介绍CRISPR/Cas系统的分类及作用机制,回顾基于CRISPR/Cas的各种生物传感器检测食源性病原体的方法,包括荧光传感器、电化学传感器和侧流层析试纸,分析应用不同Cas蛋白检测的优缺点,为食源性病原体的检测提供借鉴。     

原生质体融合结合基因编辑技术显著提高酿酒酵母2-苯乙醇产量

酿酒酵母(Saccharomyces cerevisiae)已用于生产天然2-苯乙醇(2-phenylethanol, 2-PE),但其产量低,耐受性差。该研究筛选出3株具有不同优良性状的酿酒酵母菌株,其中菌株LSC-1的2-PE产量为3.41 g/L,菌株NGER对2-PE的耐受性达3.60 g/L,菌株S.C-1耐热性好并能在41℃下生长。以这3株菌为亲本,通过2轮原生质体融合获得融合子菌株RH2-16,对2-PE的耐受性提高了20%。以L-苯丙氨酸为底物,发酵转化36 h, 2-PE产量达到4.31 g/L,与亲本菌株LSC-1和2-PE工业生产菌株CWY132相比,分别提高26.4%和38.1%。利用CRISPR/Cas9系统,在RH2-16中构建了含突变的Ras特异性鸟嘌呤核苷酸交换因子CDC25(W1416C)菌株,2-PE的产量提高了8%。在RH2-16中过表达艾氏途径合成2-PE关键基因ARO8,ARO10和ADH2未能提高2-PE产量。研究为筛选合成2-PE等天然产物的新型酿酒酵母菌株及其育种提供了一种有效策略。     

贺兰山东麓葡萄酒产区酿酒酵母的分离及其主要特性研究

为进一步丰富我国酿酒酵母资源,从贺兰山东麓葡萄酒产区的6个不同品种的成熟期酿酒葡萄果实上分离获得9株野生酿酒酵母,以商业酿酒酵母F15为对照,对分离所得菌株分别进行酒精、SO_2、葡萄糖、pH和高渗等耐受性以及β-葡萄糖苷酶活力和果糖利用率影响因素的研究。结果表明,菌株B-1比较适合用于酿造葡萄酒,且β-葡萄糖苷酶主要存在于酿酒酵母细胞外;酿酒酵母的PFK及HK的活性与果糖利用率无相关关系。     

CRISPR/Cas系统在食源性致病菌快速检测新技术中的研究进展

食源性致病菌的早期筛查与快速检测对食品安全和临床诊断至关重要。然而,传统检测方法操作繁琐、费时费力,难以满足快速检测需求。成簇、规则间隔的短回文重复序列（clustered regularly interspaced short palindromic repeats,CRISPR）和关联蛋白（CRISPR-associated protein,Cas）组成的CRISPR/Cas是广泛存在于细菌和古细菌中的一种免疫系统,其高效特异序列识别及切割活性的特性,为食源性致病菌高灵敏、快速检测提供了一种新的途径。本文介绍了CRISPR/Cas系统的原理、机制及发展,总结了近年来基于CRISPR/Cas系统结合不同结果报告方式用于食源性致病菌快速检测的最新进展,并对其优势、局限性和未来发展方向进行了讨论。     

CRISPR/Cas技术在乳酸菌中的研究进展

乳酸菌是重要的食品发酵剂,其传统的基因遗传操作技术为同源重组,该技术为后续的基因编辑工作奠定了基础,但是也存在操作繁琐、效率低等不足。成簇的规则间隔短回文重复序列(clustered regularly interspaced short palindromic repeats, CRISPR)及其相关蛋白(CRISPR-associated protein, Cas)以其高效、便捷性推动了乳酸菌基因编辑的发展。该文综述了CRISPR的原理、分类,重点阐述了CRISPR操作系统在乳酸菌方面的应用。     

CRISPR/Cas9-RNA interference system for combinatorial metabolic engineering of Saccharomyces cerevisiae

The yeast Saccharomyces cerevisiae is widely used in industrial biotechnology for the production of fuels, chemicals, food ingredients, food and beverages, and pharmaceuticals. To obtain high-performing strains for such bioprocesses, it is often necessary to test tens or even hundreds of metabolic engineering targets, preferably in combinations, to account for synergistic and antagonistic effects. Here, we present a method that allows simultaneous perturbation of multiple selected genetic targets by combining the advantage of CRISPR/Cas9, in vivo recombination, USER assembly and RNA interference. CRISPR/Cas9 introduces a double-strand break in a specific genomic region, where multiexpression constructs combined with the knockdown constructs are simultaneously integrated by homologous recombination. We show the applicability of the method by improving cis,cis-muconic acid production in S. cerevisiae through simultaneous manipulation of several metabolic engineering targets. The method can accelerate metabolic engineering efforts for the construction of future cell factories.     

CRISPR/Cpf1 enables fast and simple genome editing of Saccharomyces cerevisiae

Cpf1 represents a novel single RNA‐guided CRISPR/Cas endonuclease system suitable for genome editing with distinct features compared with Cas9. We demonstrate the functionality of three Cpf1 orthologues – Acidaminococcus spp. BV3L6 (AsCpf1), Lachnospiraceae bacterium ND2006 (LbCpf1) and Francisella novicida U112 (FnCpf1) – for genome editing of Saccharomyces cerevisiae. These Cpf1‐based systems enable fast and reliable introduction of donor DNA on the genome using a two‐plasmid‐based editing approach together with linear donor DNA. LbCpf1 and FnCpf1 displayed editing efficiencies comparable with the CRISPR/Cas9 system, whereas AsCpf1 editing efficiency was lower. Further characterization showed that AsCpf1 and LbCpf1 displayed a preference for their cognate crRNA, while FnCpf1‐mediated editing with similar efficiencies was observed using non‐cognate crRNAs of AsCpf1 and LbCpf1. In addition, multiplex genome editing using a single LbCpf1 crRNA array is shown to be functional in yeast. This work demonstrates that Cpf1 broadens the genome editing toolbox available for Saccharomyces cerevisiae. © 2017 The Authors. Yeast published by John Wiley & Sons, Ltd.     

A CRISPR/Cas9 method to generate heterozygous alleles in Saccharomyces cerevisiae

Saccharomyces cerevisiae is a genetically facile organism, yet multiple CRISPR/Cas9 techniques are widely used to edit its genome more efficiently and cost effectively than conventional methods. The absence of selective markers makes CRISPR/Cas9 editing particularly useful when making mutations within genes or regulatory sequences. Heterozygous mutations within genes frequently arise in the winners of evolution experiments. The genetic dissection of heterozygous alleles can be important to understanding gene structure and function. Unfortunately, the high efficiency of genome cutting and repair makes the introduction of heterozygous alleles by standard CRISPR/Cas9 technique impossible. To be able to quickly and reliably determine the individual phenotypes of the thousands of heterozygous mutations that can occur during directed evolutions is of particular interest to industrial strain improvement research. In this report, we describe a CRISPR/Cas9 method that introduces specific heterozygous mutations into the S. cerevisiae genome. This method relies upon creating silent point mutations in the protospacer adjacent motif site or removing the protospacer adjacent motif site entirely to stop the multiple rounds of genome editing that prevent heterozygous alleles from being generated. This technique should be able to create heterozygous alleles in other diploid yeasts and different allelic copy numbers in polyploid cells.     

New vectors for simple and streamlined CRISPR–Cas9 genome editing in Saccharomyces cerevisiae

Clustered regularly interspaced short palindromic repeats (CRISPR)–Cas9 technology is an important tool for genome editing because the Cas9 endonuclease can induce targeted DNA double‐strand breaks. Targeting of the DNA break is typically controlled by a single‐guide RNA (sgRNA), a chimeric RNA containing a structural segment important for Cas9 binding and a 20mer guide sequence that hybridizes to the genomic DNA target. Previous studies have demonstrated that CRISPR–Cas9 technology can be used for efficient, marker‐free genome editing in Saccharomyces cerevisiae. However, introducing the 20mer guide sequence into yeast sgRNA expression vectors often requires cloning procedures that are complex, time‐consuming and/or expensive. To simplify this process, we have developed a new sgRNA expression cassette with internal restriction enzyme sites that permit rapid, directional cloning of 20mer guide sequences. Here we describe a flexible set of vectors based on this design for cloning and expressing sgRNAs (and Cas9) in yeast using different selectable markers. We anticipate that the Cas9–sgRNA expression vector with the URA3 selectable marker (pML104) will be particularly useful for genome editing in yeast, since the Cas9 machinery can be easily removed by counter‐selection using 5‐fluoro‐orotic acid (5‐FOA) following successful genome editing. The availability of new vectors that simplify and streamline the technical steps required for guide sequence cloning should help accelerate the use of CRISPR–Cas9 technology in yeast genome editing. Vectors pT040, pJH001, pML104 and pML107 have been deposited at Addgene ( www.addgene.org ). Copyright © 2015 John Wiley & Sons, Ltd.     

Use of a fluoride channel as a new selection marker for fission yeast plasmids and application to fast genome editing with CRISPR/Cas9

Fission yeast is a powerful model organism that has provided insights into important cellular processes thanks to the ease of its genome editing by homologous recombination. However, creation of strains with a large number of targeted mutations or containing plasmids has been challenging because only a very small number of selection markers is available in Schizosaccharomyces pombe. In this paper, we identify two fission yeast fluoride exporter channels (Fex1p and Fex2p) and describe the development of a new strategy using Fex1p as a selection marker for transformants in rich media supplemented with fluoride. To our knowledge this is the first positive selection marker identified in S. pombe that does not use auxotrophy or drug resistance and that can be used for plasmids transformation or genomic integration in rich media. We illustrate the application of our new marker by significantly accelerating the protocol for genome edition using CRISPR/Cas9 in S. pombe. Copyright © 2016 John Wiley & Sons, Ltd.     

A history of genome editing in Saccharomyces cerevisiae

Genome editing is a form of highly precise genetic engineering which produces alterations to an organism's genome as small as a single base pair with no incidental or auxiliary modifications; this technique is crucial to the field of synthetic biology, which requires such precision in the installation of novel genetic circuits into host genomes. While a new methodology for most organisms, genome editing capabilities have been used in the budding yeast Saccharomyces cerevisiae for decades. In this review, I will present a brief history of genome editing in S. cerevisiae, discuss the current gold standard method of Cas9‐mediated genome editing, and speculate on future directions of the field.