Everything about the lab industry

When cultivating microorganisms in a laboratory, a growth environment called a culture medium is used. The culture medium may be pure chemical substances (medium with a determined chemical composition), or it may contain organic substances, or it may be composed of living organisms such as fertilized eggs. Microorganisms that grow in or on such media form cultures. If there is only one type of organism, the culture is regarded as a pure culture, and if there are populations of different organisms, it is regarded as a mixed culture. When used for the first time, the culture medium should be sterile, which means that there are no life forms before the inoculation of the microorganisms. General microbial culture medium For bacterial culture, the commonly used medium is nutrient broth, which is a liquid containing protein, salt and growth promoters, which can support many bacteria. In order to solidify the medium, add agar and other reagents. Agar is a polysaccharide, it does not add nutrients to the medium, but only solidifies it. The resulting medium is nutrient agar. Many media for microorganisms are complex and reflect the growth requirements of microorganisms. For example, most fungi require additional carbohydrates and an acidic environment to achieve optimal growth. The medium used for these organisms is potato dextrose agar, also known as Sabouraud dextrose agar. For protozoa, liquid culture medium is generally required. For rickettsiae and virus, living tissue cells must be provided for optimal culture. For anaerobic microorganisms, the atmosphere must be anaerobic. To eliminate oxygen, the culture medium can be placed in a container that produces carbon dioxide and hydrogen and removes oxygen from the atmosphere. Commercial products meet these conditions. Anaerobic chambers can also be used in closed compartments where technicians can manipulate the culture medium. In order to promote the formation of carbon dioxide, candles can be burned to consume oxygen and replaced with carbon dioxide. Special microbial culture medium Certain microorganisms are cultivated in selective media. These media prevent the growth of unwanted organisms while promoting the growth of desired organisms. For example, mannitol salt agar is selective for staphylococci because most other bacteria cannot grow in its high salt environment. Another selective medium is Bright Green Agar, which inhibits Gram-positive bacteria while allowing the growth of Gram-negative bacteria such as Salmonella. There are other cultural media that are different media. These media provide an environment in which different bacteria can be distinguished. For example, Fuchsia Bile Agar is used to distinguish coliforms, such as Escherichia coli and non-coliforms. The coliforms are bright pink colonies in this medium, while the non-coliforms are pale pink or transparent. Certain media are both selective and differentiated. For example, MacConkey Agar distinguishes lactose-fermenting bacteria from non-lactose-fermenting bacteria, while inhibiting the growth of gram-positive bacteria. Since lactose-fermenting bacteria are often associated with water pollution, they can be distinguished by adding a water sample to MacConkey agar and waiting for growth to appear. In some cases, it is necessary to prepare enriched media. This medium provides specific nutrients and can promote the reproduction of specific types of microorganisms in mixed samples. For example, when trying to isolate Salmonella from a stool sample, it is helpful to place the material sample in an enriched medium to promote the propagation of Salmonella before the isolation technique begins. In order to use microorganisms in the laboratory, they need to be obtained in pure culture. A pure culture of bacteria can be obtained by spreading the bacteria and allowing individual cells to form growing clumps called colonies. You can then pick a sample from the colony and make sure it contains only one type of bacteria. Culturing these bacteria on a separate medium will produce a pure culture. In order to preserve microbial cultures, they can be placed in the refrigerator to slow down the rate of metabolism. The other two methods are deep freezing and freeze drying. For deep freezing, microorganisms are placed in a liquid and quickly frozen at a temperature below –50°C. Freeze-drying (lyophilization) is carried out in equipment that uses vacuum pumping after freezing the microbial suspension. The culture is similar to a powder, and the microorganisms can be stored for a long time under this condition.

Successful recombinant protein expression and purification are often essential for basic research as well as biotechnology and commercial applications. High-throughput protein expression and purification in Escherichia coli has begun to revolutionize the research methods in various research fields. Experiments that are usually performed manually are processing one protein at a time in a few weeks, and now hundreds of proteins can be performed in just one week. However, limitations still exist, and further improvements are possible. The ease of genetic manipulation, low cost, rapid growth and the number of previous studies have made E. coli one of the most widely used microbial species for the production of recombinant proteins. In this post-genomic era, rapid expression and purification of large numbers of proteins for academic and commercial purposes in a high-throughput manner is still facing challenges. At present, several state-of-the-art methods are suitable for the cloning, expression and purification of a large number of molecules, and the latest developments related to soluble protein expression, mRNA folding, fusion tags, post-translational modifications and membrane protein production are discussed. HTP High-throughput research can be defined as research that allows the measurement results of thousands of biomolecules to be obtained at the same time, thus making large-scale repetitions possible. In the post-genomic era, the use of high-throughput technologies for measuring DNA, RNA, proteins, lipids, and metabolites has increased dramatically. These technologies have been successfully applied to answer various biological questions related to cancer biology. Protein expression and purification play a central role in biochemistry. Recombinant proteins can be expressed using prokaryotic systems (E. coli and Bacillus subtilis), eukaryotic systems (yeast, insect cells and mammalian cells) or in vitro systems. The E. coli system is the host of choice for preliminary screening of recombinant protein expression, because these cells can be easily manipulated, cultivated inexpensively, and grow rapidly. In recent years, many new strains, vectors, and tags have been developed to overcome the limitations of the system, including codon bias, inclusion body formation, toxicity, protein inactivation, mRNA instability, and lack of post-translational modifications. It has been extensively studied in the E. coli expression system, but it is labor-intensive and time-consuming to use this system for protein expression and purification. Therefore, parallel, high-throughput methods must be used for protein expression and purification, which has always been the bottleneck of protein function, structure and application research in the post-genomic era. High-throughput protein production methods are widely used, and recombinant proteins in the form of inclusion bodies are even expressed and purified in a parallel manner. Effective promoter An effective promoter for expressing heterologous proteins in E. coli has four key features: First, the promoter is strong enough to allow the accumulation of recombinant protein to be greater than or equal to 10-30% of the total cell protein; second, it shows the minimum basic transcriptional activity, so unnecessary transcription is avoided before induction; third, the promoter can be induced simply and cheaply; fourth, the promoter activity can be precisely adjusted. The selection of strains for expressing recombinant proteins also plays an important role in protein expression, solubility and yield. A few E. coli strains such as BL21 and its derivatives are widely used. Different E. coli strains promote the expression of proteins containing disulfide bonds or proteins encoded by genes that contain rare codons and proteins that are toxic to E. coli. In addition, co-expression with some genes increased the expression of post-translationally modified proteins. So far, several E. coli strains that can significantly increase membrane protein production have been designed

Most human genomes can be transcribed into RNA, but only a few regions produce protein-coding mRNA, while the remaining regions are transcribed into non-coding RNA. Long non-coding RNAs (lncRNAs) are known for their important regulatory roles in a variety of biological processes, such as imprinting, dose compensation, transcriptional regulation, and splicing. In the past 30 years, through genome editing of pluripotent stem cells (PSC), the physiological functions of protein-coding genes have been extensively characterized. However, the research on lncRNAs using genome editing technology has only received attention. In recent years, thousands of lncRNAs have been identified. Some of them have proven to play an important role in PSC. However, advances in genome editing technology have only just begun to be widely used in PSC to study the function of lncRNA. Considering the different functions of lncRNA genomic loci and their transcripts, a variety of genome editing methods should be used to distinguish the functions of lncRNA transcripts and their loci in PSC. lncRNAs are important biomarkers of embryonic development and disease progression. The establishment of the lncRNA reporter gene in the body will enable the monitoring of these processes. The development of emerging CRISPR genome editing technology has opened new doors for lncRNA biology in PSC. Future research should adopt these new strategies to explore the function of lncRNA in PSC. These genome editing tools should also be used to explore the physiological functions of lncRNA in the system. lncRNA knockout Molecular scissors have been used to generate point mutations in key domains of protein-coding genes, which in turn induce premature termination of translation to knock out genes. Unlike protein-coding genes, the transcriptional functional domains of most lncRNAs are still unclear; therefore, it is impossible to study lncRNAs through loss-of-function mutagenesis. Therefore, to knock out lncRNA, it is necessary to completely or partially delete the lncRNA gene. In order to avoid the indirect effects of lncRNA knockout, researchers need to manipulate lncRNA genomic loci without affecting the genomic characteristics of other genes. This, however, is difficult to achieve in some cases. For lncRNAs located in the promoter regions of other genes or overlapping with the exons of protein-coding genes, partial deletion of them through genome editing can only be applied when the expression of other genes is not affected. For the lncRNA in the intron of the protein-coding gene, it is necessary to delete the lncRNA gene without interfering with the splicing of the intron region. The lncRNA in the intergenic region is far away from other genes and can be easily removed by genome editing technology in a manner similar to protein-coding genes. However, intergenic lncRNA loci that overlap with enhancers, such as enhancer RNA, are also difficult to study with genome editing, because the deletion of these sites may interfere with enhancer function and affect the expression of distant genes. With the discovery of more lncRNAs and the further understanding of the function of lncRNAs, the knockout of lncRNAs based on homologous recombination is more widely used to delete lncRNAs in PSCs. In one of the studies, 18 lncRNA genes in mouse ESC were knocked out to produce lncRNA knockout mice. Replacing the lncRNA locus with the lacZ reporter gene allows visualization of the spatiotemporal expression patterns of these lncRNAs in animal models. Another similar study created knockout ESC lines for 20 lncRNAs through gene targeting, and used these ESCs to create knockout mice to study the broad role of lncRNA in mice. These lncRNA knockout mice constitute a valuable supplement to resources for studying the physiological role of lncRNA. Genome editing strategies A single method is not enough to identify all possible functions of lncRNA. A variety of genome editing strategies need to be adopted to discover the function of lncRNA and its transcription site in PSC. Since CRISPR/Cas9-based genome editing technology can effectively delete large fragments of the genome, it has been used to perform genome-wide screening of lncRNA functions. Use multiple gRNAs against a single lncRNA to achieve effective ablation of lncRNA gene expression; therefore, the paired gRNA library can only target hundreds of lncRNAs. Using this method, lncRNAs, which are critical to the survival of cancer cells, have been identified. Another way to use CRISPR-Cas9 to regulate lncRNA is through CRISPR interference (CRISPRi), which is a fusion of inactivated Cas9 (dCas9) and transcription inhibitors (such as KRAB). By recruiting dCas9-KRAB to the promoter region of lncRNA with multiple gRNAs, the expression of lncRNA is hindered by transcription repressors recruited by KRAB protein. CRISPRi was used to manipulate the expression of lncRNA (GAS5, H19, MALAT1, NEAT1, TERC and XIST) in K562 cells.

Ion channels are involved in various basic physiological processes, and their failure can lead to a variety of human diseases. Therefore, ion channels represent an attractive class of drug targets and an important class of off-target proteins, which can be used for in vitro pharmacological analysis. Over the past few decades, rapid advances in functional assays and instrument development have enabled high-throughput screening (HTS) activities in an ever-expanding list of channel types. In chronological order, HTS methods for ion channels include ligand binding assays, flux-based assays, fluorescence-based assays, and automated electrophysiological assays. Ligand binding assays Ligand binding assays have been widely used to screen ion channel modulators. However, these assays are not considered functional assays because they measure the binding affinity of the compound to the ion channel, rather than the ability to change the channel's function. Ligand binding assays require prior knowledge of target binding sites and the formation of radiolabeled ligands specific to these binding sites. The activity of the test compound is indicated by the displacement of the labeled ligand. Therefore, conventional instruments can be used, where throughput represents its main advantage. Because this method only finds that the compound affects radioligand binding, it ignores allosteric modulators of ion channels. The binding assay can identify affinity data, but cannot identify changes in ion channel function. Ion flux Ion flux measurement has been successfully applied to directly access functional changes in ion channel activity. Radioisotopes have been used to track the cell inflow or outflow of specific ions (such as 22 Na +, 45 Ca 2+ and 86 Rb +), and are used to study Na +, Ca 2+ and K + channels, respectively. The commonly used measurement format is 86 Rb + efflux of K + channel or non-selective cation channel. This detection technology is widely used in the pharmaceutical industry for two drug discovery and hERG-related drug safety screening to determine potential QT liabilities that may cause fatal arrhythmia. Fluorescenc Fluorescence-based methods cannot directly measure ion currents. They measure membrane potential-dependent or ion-concentration-dependent changes in fluorescence signals due to ion flux. Because fluorescence-based methods can produce reliable and uniform cell population measurements, these assays are similar to those of other protein classes. Therefore, more instrument options and expertise can be applied. These assays are relatively easy to implement and optimize to achieve higher throughput. Patch clamp Patch clamp has been widely regarded as the gold standard for direct recording of ion channel activity. This technology can provide high-quality and physiologically relevant data of ion channel functions at the single cell or single channel (in a small piece of membrane) level. For pharmacological testing of compounds, it provides a standard for measuring the potency of the interaction between the compound and the channel. Although the conventional patch clamp provides a direct, information-rich real-time method to study channel functions, it has very low throughput and labor-intensive characteristics. Combination At present, the combination of fluorescence-based screening technology and automatic patch clamp has become the most commonly used method for ion channel targeted drug discovery. As costs decrease and technology advances, automated electrophysiology will become the main form of measurement for most ion channel subtypes. For different ion channel subclasses, high-throughput screening methods differ due to ion selectivity, channel activation kinetics, and consideration of whether ligands are required. Advances and improvements in ion channel HTS technology have accelerated the discovery of ion channel drugs. These measurements have relatively low time resolution and information content, but can be robust and low-cost. The electrophysiological method has the most direct method to measure ion channel activity and also provides flexibility for the analysis and optimization of each channel type. The combination of non-electrophysiological and electrophysiological HTS methods provides an integrated and cost-effective method for ion channel drug discovery and ensures the generation of high-quality data.