2811 月/24
Macrophage Immortalization

KMD Bioscience-Macrophage Immortalization

CRO Service

Immortalization of macrophages is particularly challenging due to their terminally differentiated state and specialized immune functions. However, several methods have been developed to immortalize macrophages to allow their long-term culture and use in research. Immortalized macrophage cell lines provide a valuable tool for studying immune responses, pathogen interactions, and inflammatory processes, offering a consistent and reproducible model for experiments.

Methods of Macrophage Immortalization

SV40 Large T Antigen

Mechanism

SV40 large T antigen immortalizes macrophages by inactivating the tumor suppressor proteins p53 and pRb, which normally regulate cell cycle progression and apoptosis. By inhibiting these proteins, SV40 large T antigen allows macrophages to bypass senescence and proliferate indefinitely.

Examples

J774A.1: A mouse macrophage cell line derived from a reticulum cell sarcoma that has been immortalized using SV40 large T antigen. J774A.1 cells are commonly used for studying phagocytosis, cytokine production, and macrophage responses to pathogens.

RAW 264.7: Although not directly immortalized by SV40, this mouse macrophage-like cell line is widely used in immune research, especially for studying macrophage activation, cytokine production, and inflammatory responses.

v-myc/v-raf Oncogenes (Bone Marrow-Derived Macrophages Immortalization)

Mechanism: The v-myc and v-raf oncogenes are used to immortalize bone marrow-derived macrophages (BMDMs). The v-myc oncogene promotes cell proliferation, while v-raf provides additional signaling to prevent apoptosis. Together, these oncogenes allow macrophages to proliferate while maintaining many of their functional characteristics.

Examples

Bac1.2F5: A murine macrophage cell line immortalized using v-myc and v-raf oncogenes. These cells retain many functional properties of primary macrophages, including the ability to produce cytokines, phagocytose, and respond to pathogens.

MPI Cells: These are murine bone marrow-derived macrophages immortalized using a combination of v-myc and v-raf oncogenes. MPI cells are used for studies on macrophage differentiation and function.

Human Telomerase Reverse Transcriptase (hTERT)

 Mechanism

Overexpression of hTERT in macrophages prevents telomere shortening, a key mechanism of cellular aging. By maintaining telomere length, hTERT immortalization enables macrophages to bypass senescence and continue dividing.

Advantages

This method is less likely to induce genetic instability compared to viral oncogenes or antigens, and cells typically retain more of their normal physiological functions.

 Challenges

Macrophages are terminally differentiated cells, which makes this method more challenging and less commonly used than in other cell types. However, it has potential for extending the life span of macrophage precursor cells (e.g., monocytes).

Retroviral Transduction with Oncogenes (e.g., c-myc, raf/ER, etc.)

Mechanism

Retroviruses can be used to introduce oncogenes such as c-myc, which promotes cell proliferation, and an estrogen receptor-regulated form of the raf oncogene (raf/ER). This method can be applied to macrophage precursor cells, such as monocytes, to allow continuous proliferation while preserving differentiation capacity.

Examples

THP-1: A human monocytic cell line derived from a leukemia patient. THP-1 cells can be differentiated into macrophage-like cells using phorbol 12-myristate 13-acetate (PMA). While THP-1 cells are not immortalized by c-myc or raf, they serve as an important model for studying human macrophage differentiation and function.

U937: A human monocyte-like cell line derived from histiocytic lymphoma. U937 cells can also be differentiated into macrophage-like cells and are used extensively in immunology research.

CRISPR/Cas9-Mediated Knockout of Tumor Suppressor Genes

Mechanism

CRISPR/Cas9 gene-editing technology can be used to knock out tumor suppressor genes, such as p53 or p16, allowing macrophages or their precursors to bypass normal cellular senescence. This technique is still under development for macrophage immortalization, but it holds promise for creating macrophage cell lines with specific genetic modifications.

 Advantages

Allows for precise gene editing and control over the immortalization process.

Challenges

This method is technically complex and may not always result in functional macrophage-like cells, as macrophages are terminally differentiated.

Conditional Immortalization Using Temperature-Sensitive SV40 T Antigen

 Mechanism

In this approach, macrophages are immortalized using a temperature-sensitive variant of SV40 large T antigen. The cells proliferate at the permissive temperature (33°C), but they can revert to a more differentiated and functional state when shifted to the non-permissive temperature (37°C). This allows researchers to maintain immortalized cells that can still differentiate and function under certain conditions.

Examples

Immortalized Bone Marrow-Derived Macrophages (iBMDM): These are murine macrophage precursors immortalized with a temperature-sensitive SV40 T antigen. They proliferate at lower temperatures and differentiate into macrophage-like cells at physiological temperatures.

 Key Considerations for Macrophage Immortalization

Retention of Functionality

Immortalized macrophages must retain critical macrophage functions, such as phagocytosis, cytokine production, and antigen presentation. Some immortalization techniques can alter cell behavior, so it is essential to validate that the cells still exhibit macrophage-like characteristics.

Genetic Stability

Some immortalization methods, particularly those involving viral oncogenes or SV40 large T antigen, may induce genetic instability or transformation, which can lead to unwanted changes in cell behavior over time. Researchers must regularly check for potential chromosomal alterations or mutations in immortalized macrophage lines.

Comparison to Primary Macrophages

While immortalized macrophage cell lines offer the advantage of unlimited proliferation, it is important to compare the behavior and responses of immortalized lines to primary macrophages to ensure the validity of experimental findings.

Species Specificity

Many immortalized macrophage cell lines (e.g., RAW 264.7, J774A.1) are derived from mice. While these lines are valuable for many studies, researchers must be cautious when extrapolating findings to human biology, as there can be differences between human and murine macrophages.

 Applications of Immortalized Macrophage Cell Lines

Immunology Research: Studying immune responses, cytokine production, and interactions with pathogens such as bacteria, viruses, and fungi.

Inflammation Studies: Investigating the role of macrophages in chronic inflammatory conditions, such as rheumatoid arthritis, atherosclerosis, and obesity.

Drug Screening: Testing the effects of immunomodulatory drugs, anti-inflammatory agents, and antimicrobial compounds on macrophage function.

Cancer Research: Studying the role of macrophages in tumor microenvironments and their contribution to cancer progression and metastasis.

Host-Pathogen Interactions: Exploring how macrophages recognize, engulf, and destroy pathogens, and how pathogens evade the immune response.

In conclusion, immortalized macrophage cell lines are invaluable tools for studying macrophage biology in a controlled and reproducible manner. Several methods, such as SV40 large T antigen, oncogene expression, and temperature-sensitive variants, allow for the creation of cell lines that can proliferate indefinitely while retaining many functional properties of primary macrophages. These immortalized cell lines are essential for advancing our understanding of immune responses, inflammatory processes, and macrophage-related diseases.

2711 月/24
b cell immortalization

KMD Bioscience-B Cell Immortalization

CRO Service

B cell immortalization is a process used to generate B cells that can proliferate indefinitely in culture, enabling continuous production of antibodies or facilitating research on B cell biology. Normally, B cells are short-lived after activation, but immortalization techniques allow them to bypass cellular senescence and keep dividing. Immortalized B cells are widely used for producing monoclonal antibodies, studying immune responses, and understanding B cell-related diseases such as lymphomas.

Key Methods of B Cell Immortalization

Epstein-Barr Virus (EBV) Transformation

Mechanism

The Epstein-Barr virus (EBV) is a herpesvirus that can infect B cells and induce their immortalization. EBV binds to the CD21 receptor on B cells, leading to the activation of signaling pathways that promote continuous cell division.

Advantages

It maintains immunoglobulin (antibody) production, making it useful for generating monoclonal antibodies.

The technique is relatively simple and widely used.

Disadvantages

The B cells may have limited proliferative capacity after a few months.

The transformation can result in non-specific activation of B cells, leading to a mix of antibody production.

Fusion with Myeloma Cells (Hybridoma Technology)

Mechanism

This is the classical method for generating monoclonal antibodies, pioneered by Köhler and Milstein in 1975. It involves fusing an antibody-producing B cell with a myeloma (cancerous plasma) cell to create a hybridoma cell line.

Advantages

Hybridoma cells proliferate indefinitely and produce a specific monoclonal antibody.

This method remains the gold standard for producing monoclonal antibodies.

Disadvantages

The process can be labor-intensive and technically demanding.

Not all B cells will successfully fuse, and careful selection is required to isolate the hybridoma cells producing the desired antibody.

Transgenic Expression of Oncogenes

Mechanism: B cells can be immortalized by introducing oncogenes, such as Bcl-6 or Myc, through viral vectors or transfection. These genes regulate cell survival and proliferation, enabling the B cells to divide indefinitely.

Advantages

Allows for controlled genetic manipulation of B cells.

Can be used to create long-term B cell lines for specific research purposes.

Disadvantages

Genetic manipulation may alter the natural behavior and function of the B cells, affecting their use in some applications.

CRISPR/Cas9-based Genetic Manipulation

Mechanism: CRISPR/Cas9 technology can be used to knock out genes responsible for senescence or introduce genes promoting proliferation, effectively immortalizing B cells.

Advantages

Offers precise and targeted manipulation of B cell genetics.

Can be used to study specific gene functions in B cell development or disease.

Disadvantages

The immortalized B cells may not behave exactly like normal B cells.

It is a relatively new method and requires technical expertise.

Applications of Immortalized B Cells

Monoclonal Antibody Production

Immortalized B cells, especially hybridomas, are widely used to produce monoclonal antibodies. These antibodies are essential tools in diagnostics, therapeutics (e.g., cancer treatment, autoimmune diseases), and research.

Study of B Cell Biology

Immortalized B cells provide a model to study B cell development, activation, differentiation, and antibody production in vitro over extended periods.

Disease Research

B cell immortalization is useful for studying diseases related to B cell dysfunction, such as B cell lymphomas, leukemias, and autoimmune diseases (e.g., lupus, rheumatoid arthritis).

Vaccine Development

Immortalized B cells are used to develop vaccines by producing specific antibodies and understanding the immune response to pathogens.

Cancer Research

Immortalized B cells can be studied to explore mechanisms behind B cell cancers like multiple myeloma, Burkitt’s lymphoma, and chronic lymphocytic leukemia (CLL).

Advantages of B Cell Immortalization

Continuous Growth: The cells proliferate indefinitely, eliminating the need to continuously source new B cells from donors or animals.

Antibody Production: Immortalized B cells (especially hybridomas) are a reliable source of specific monoclonal antibodies.

Consistency: Immortalized cells provide a consistent and reproducible model for experiments and antibody production.

 Limitations of B Cell Immortalization

Genetic Changes: Immortalized cells may undergo genetic alterations that change their behavior compared to primary B cells.

Functional Differences: The immortalized B cells may not fully represent normal B cell physiology and may have altered responses to stimuli.

Virus Integration: In EBV-immortalized cells, viral gene expression may affect cell behavior and complicate some types of research.

In summary, B cell immortalization is a powerful tool in immunology and biotechnology, offering sustained B cell function for antibody production, disease research, and drug development. Various techniques like EBV transformation, hybridoma technology, and genetic manipulation enable researchers to explore and harness B cell functions for scientific and therapeutic purposes.

2611 月/24
cell immortalization

KMD Bioscience-Cell Immortalization

CRO Service

Cell immortalization refers to the process of modifying normal cells to allow them to proliferate indefinitely. Under natural circumstances, most cells have a limited ability to divide due to a phenomenon known as replicative senescence, which occurs as a result of telomere shortening and other cellular stress factors. Immortalization overcomes this limitation by enabling cells to bypass senescence and avoid programmed cell death (apoptosis), allowing them to grow continuously.

Immortalized cells are widely used in research because they provide a consistent, long-term source of cells for experiments, such as studying cellular functions, drug screening, cancer biology, and genetic studies.

Mechanisms of Cell Immortalization

There are several methods to immortalize cells, typically targeting pathways that control senescence and apoptosis. These methods involve altering genes or introducing factors that enable the cells to divide indefinitely.

Telomerase Activation

Telomeres are repetitive nucleotide sequences at the ends of chromosomes that protect them from damage. With each cell division, telomeres shorten, eventually leading to replicative senescence. This mechanism limits the number of divisions a cell can undergo, a phenomenon known as the Hayflick limit.

Telomerase is an enzyme that elongates telomeres, allowing cells to bypass senescence. In most somatic cells, telomerase is inactive, but it is highly active in certain cell types (e.g., germ cells, stem cells, and cancer cells).

Telomerase reverse transcriptase (TERT), the catalytic subunit of telomerase, is commonly introduced into cells to reactivate telomerase and maintain telomere length, which extends the replicative lifespan of the cells. This method is often used to immortalize human fibroblasts and epithelial cells.

Viral Oncogenes (SV40 Large T Antigen)

One of the most common methods of immortalizing cells is by introducing viral oncogenes, such as SV40 large T antigen, derived from Simian virus 40 (SV40). This viral protein disrupts key tumor suppressor pathways, particularly those regulated by the p53 and Rb (retinoblastoma) proteins.

p53: A tumor suppressor that regulates cell cycle arrest and apoptosis in response to DNA damage or stress. SV40 large T antigen binds and inactivates p53, preventing apoptosis and allowing cells to continue dividing.

Rb (Retinoblastoma protein): Rb regulates the G1-S transition in the cell cycle. By binding to and inactivating Rb, SV40 large T antigen allows cells to bypass cell cycle checkpoints, leading to continuous proliferation.

SV40 large T antigen is widely used to immortalize a variety of cell types, including fibroblasts, epithelial cells, and neurons.

Human Papillomavirus (HPV) Oncoproteins (E6 and E7)

Human papillomavirus (HPV), particularly HPV16 and HPV18, contains two key oncogenes, E6 and E7, which can immortalize cells by interfering with the p53 and Rb pathways, similar to SV40 large T antigen.

E6: Promotes the degradation of p53, preventing cell cycle arrest and apoptosis.

E7: Binds and inactivates Rb, allowing cells to bypass the G1 checkpoint and proliferate.

HPV E6 and E7 are commonly used to immortalize human epithelial cells, such as keratinocytes and cervical cells.

hTERT Overexpression

As mentioned earlier, the human telomerase reverse transcriptase (hTERT) gene can be introduced into cells to maintain telomere length, preventing replicative senescence. This is a particularly clean method for immortalization because it does not affect the normal cell cycle control pathways (unlike SV40 T antigen or HPV E6/E7).

hTERT immortalization is widely used for human fibroblasts, epithelial cells, and endothelial cells, and it is preferred when studying cellular behavior without disrupting tumor suppressor pathways.

Bypass of the CDK Inhibitors

Cyclin-dependent kinase inhibitors (CDKIs), such as p16^INK4a, regulate the cell cycle by inhibiting cyclin-dependent kinases (CDKs) that control the progression from G1 to S phase.

In some immortalization methods, particularly in mouse cells, knocking down or silencing p16^INK4a (or its human equivalent p21^CIP1) can prevent cells from entering senescence and allow them to continue dividing.

Applications of Immortalized Cells

Basic Research

Immortalized cell lines provide a consistent and reproducible cell source for studying cellular functions, signaling pathways, and gene expression.

Examples of immortalized cell lines include HeLa cells (derived from cervical cancer), HEK293 cells (human embryonic kidney cells), and 3T3 fibroblasts (mouse fibroblasts).

Cancer Research

Since many cancer cells acquire immortality through similar mechanisms (e.g., reactivation of telomerase or p53 inactivation), immortalized cell lines serve as models for studying tumor biology, cancer progression, and the effects of anti-cancer drugs.

Drug Screening and Development

Immortalized cells are widely used in high-throughput screening assays to test the efficacy and toxicity of new drug candidates.

They provide a reproducible and scalable platform for screening large compound libraries in pharmaceutical research.

Gene Editing and Functional Studies

Immortalized cells are used in CRISPR-Cas9 and other gene-editing techniques to study the roles of specific genes in cellular processes such as proliferation, differentiation, and apoptosis.

Biomanufacturing

Immortalized cells can be engineered to produce therapeutic proteins, antibodies, or vaccines. Cell lines like Chinese hamster ovary (CHO) cells are widely used in the biotechnology industry for producing biopharmaceuticals.

Vaccine Production

Immortalized cell lines are commonly used in vaccine production, as they can grow rapidly and provide a reliable platform for propagating viruses or producing viral antigens.

Vero cells (derived from African green monkey kidney cells) are an example of an immortalized cell line used for vaccine production.

 Examples of Commonly Used Immortalized Cell Lines

HeLa Cells

Derived from cervical cancer, HeLa cells are one of the first and most widely used human cell lines in research. They were immortalized naturally by the HPV E6 and E7 oncoproteins.

HEK293 Cells

Human embryonic kidney cells immortalized with adenovirus E1A/E1B oncogenes. Widely used for gene expression, protein production, and drug screening.

NIH 3T3 Cells

Mouse embryonic fibroblasts that were spontaneously immortalized through serial passaging. Commonly used in molecular biology and cancer research.

CHO Cells

Chinese hamster ovary cells are used extensively in biomanufacturing for producing recombinant proteins, monoclonal antibodies, and vaccines.

293T Cells

A derivative of HEK293 cells, modified with SV40 large T antigen for enhanced gene expression. Frequently used for transfection and viral production.

Vero Cells

Derived from the kidney of an African green monkey, Vero cells are commonly used for virology research and vaccine production.

Challenges and Considerations

Genetic Stability

Immortalized cell lines can undergo genetic changes over time, leading to variability in behavior. This can be problematic for reproducibility across experiments.

Loss of Normal Cellular Phenotypes

While immortalization allows cells to proliferate indefinitely, it can also alter normal cell cycle control, differentiation capacity, or other phenotypic characteristics. Researchers need to validate that key functions of interest are preserved after immortalization.

Risk of Transformation

Some immortalization methods, particularly those using viral oncogenes (e.g., SV40 large T antigen or HPV E6/E7), can increase the risk of cell transformation and tumorigenicity. This should be considered when choosing the appropriate immortalization strategy.

Conclusion

Cell immortalization is a valuable technique for generating cell lines that can proliferate indefinitely, providing an unlimited source of cells for research, drug development, and biomanufacturing. Various methods, including telomerase activation, viral oncogene expression, and CDK inhibitor bypass, are used depending on the cell type and research goals. While immortalized cell lines offer many advantages, they must be carefully characterized to ensure they retain relevant biological properties for specific applications.

2211 月/24

KMD Bioscience-Yeast Display Sorting

Antibody Discovery

Yeast display sorting is a critical step in the yeast surface display process, used to identify and isolate yeast cells displaying peptides or proteins with desirable binding properties, such as high affinity or specificity for a target molecule. This process relies heavily on fluorescence-activated cell sorting (FACS), which allows researchers to quantitatively measure the binding of displayed proteins or peptides to fluorescently labeled targets and then isolate the cells that exhibit the strongest binding.

 Overview of Yeast Display Sorting Process

Expression of Protein/Peptide Libraries on Yeast Surface

The yeast cells display a library of protein or peptide variants on their surface, typically fused to a surface anchor protein like Aga2p. These variants could be antibodies, enzymes, peptides, or other protein fragments.

 Binding to Target Molecule

Yeast cells are incubated with a fluorescently labeled target (e.g., a protein, receptor, antigen, or small molecule). Yeast cells displaying peptides or proteins that bind to the target will retain the fluorescent label on their surface.

Fluorescence-Activated Cell Sorting (FACS)

After incubation with the labeled target, the yeast cells are passed through a flow cytometer, where cells are sorted based on their level of fluorescence. Cells displaying proteins or peptides that bind strongly to the target exhibit higher fluorescence intensity and are separated from those with weaker or no binding.

Selection and Enrichment of High-Affinity Binders

The yeast cells with the highest fluorescence intensity (i.e., those displaying the best binders) are collected and cultured. This population can then undergo multiple rounds of sorting to further enrich for variants with even higher affinity or specificity.

 Steps in the Yeast Display Sorting Process

Preparation of Yeast Cells Displaying Protein/Peptide

Induce Surface Display: Yeast cells containing the library are cultured under conditions that induce the expression of the gene encoding the protein or peptide of interest fused to the Aga2p surface anchor. This results in the display of these proteins/peptides on the surface of the yeast cells.

Incubation with Labeled Target

The yeast cells are incubated with a fluorescently labeled target. The target can be labeled with a fluorescent dye such as FITC, PE, or Alexa Fluor. If the protein or peptide displayed on the yeast surface binds to the target, the yeast cells will be fluorescently labeled.

Secondary Labeling (Optional): In some cases, a secondary fluorescent antibody may be used to detect the protein of interest or specific epitope tags like FLAG or HA, to confirm surface display of the fusion protein.

Fluorescence-Activated Cell Sorting (FACS)

Sorting Based on Fluorescence:

The yeast cells are passed through a flow cytometer, where each cell’s fluorescence is measured as it flows through a laser beam. The cells are sorted into different populations based on the intensity of the fluorescence signal.

Dual-Color Sorting: Often, two fluorescent signals are measured:

Protein Display: One fluorescent signal is used to confirm that the protein or peptide is being properly displayed on the yeast surface (e.g., through an epitope tag like HA or FLAG).

Target Binding: The other fluorescent signal is used to measure the binding of the displayed protein or peptide to the fluorescently labeled target.

Gating for Selection

Researchers set a gate to select yeast cells that show both high levels of surface protein display and strong binding to the target. These cells are separated from the rest of the population and collected for further rounds of sorting or analysis.

Negative Sorting (Optional): If specificity is a concern, yeast cells can also be incubated with an irrelevant target or labeled non-target molecule to eliminate non-specific binders. Cells that bind to the non-target molecule are excluded from the final selection.

Enrichment of High-Affinity Binders

Multiple Rounds of Sorting:

The yeast cells collected after the first round of sorting can be expanded in culture and subjected to additional rounds of sorting. Each round refines the population, enriching for yeast cells displaying peptides or proteins with higher affinity or specificity for the target.

Typically, 2–4 rounds of sorting are performed, with progressively narrower gates in FACS to select the highest affinity binders.

Analysis and Recovery of Selected Clones

Sequencing of Selected Variants:

After sorting, the DNA encoding the protein or peptide from the selected yeast clones is recovered and sequenced to identify the sequences of the high-affinity binders.

Next-Generation Sequencing (NGS): NGS can be used to analyze the entire population of binders, allowing for detailed insights into the diversity of selected variants and the frequency of specific mutations.

Functional Validation:

The selected clones can be subjected to further biochemical and functional assays to validate the binding affinity and specificity. Common methods include surface plasmon resonance (SPR), enzyme-linked immunosorbent assays (ELISA), and competition binding assays.

 Applications of Yeast Display Sorting

Antibody Engineering and Selection

Yeast display sorting is widely used in antibody discovery and engineering to identify antibody fragments (such as scFvs or Fabs) with high affinity for antigens. By sorting libraries of antibody variants, researchers can isolate clones that bind tightly to a target antigen.

Protein-Protein Interaction Studies

Yeast display sorting is used to study protein-protein interactions by screening peptide or protein libraries for variants that bind specifically to a target protein.

Directed Evolution and Protein Engineering

Through iterative rounds of yeast display sorting, proteins can be evolved to improve properties like binding affinity, stability, or enzymatic activity. This method is frequently applied to optimize therapeutic proteins, enzymes, and receptors.

Peptide Display and Screening

Peptide libraries displayed on yeast can be sorted to identify binding peptides that interact with a target protein, receptor, or other molecules. These peptides can be used in therapeutic development or as molecular tools to study biological pathways.

Epitope Mapping

Yeast display sorting can be used to identify the specific regions (epitopes) of a protein that are recognized by antibodies. By displaying fragments or peptides of the target protein on yeast and sorting based on antibody binding, epitope mapping can be performed.

Drug Discovery

Yeast display libraries can be used to screen for small peptides or protein domains that bind to drug targets, providing potential leads for therapeutic development.

 Advantages of Yeast Display Sorting

Quantitative Analysis: FACS allows for precise, quantitative measurement of binding interactions, enabling the selection of the best binders with the highest affinity.

High-Throughput Screening: Yeast display sorting enables the screening of large libraries of protein or peptide variants in a high-throughput manner, making it ideal for directed evolution and protein engineering.

Eukaryotic System: Yeast cells provide a eukaryotic environment for protein folding and post-translational modifications, making yeast display more biologically relevant for certain proteins than bacterial display systems.

Multiple Rounds of Enrichment: The ability to perform multiple rounds of sorting allows for the progressive enrichment of clones with higher affinity or specificity, leading to the selection of the best possible variants.

 Challenges and Limitations of Yeast Display Sorting

Glycosylation Differences: Yeast glycosylation patterns differ from those of mammalian cells, which may affect the binding or activity of certain glycosylated proteins.

Size Limitations: Larger proteins or complex protein structures may not display efficiently on the yeast surface, which can limit the types of proteins that can be screened using this method.

Off-Target Binding: Non-specific binding can sometimes occur, especially when the target protein is present in high concentrations. Negative sorting steps can mitigate this, but careful optimization is needed.

 Conclusion

Yeast display sorting, particularly using FACS, is a powerful tool for identifying high-affinity protein and peptide binders from large libraries. By enabling high-throughput and quantitative screening, this method has revolutionized protein engineering, antibody discovery, and functional genomics. Through multiple rounds of sorting, yeast display allows for the enrichment of clones with optimal binding properties, making it a critical method for directed evolution and drug discovery.

2111 月/24

KMD Bioscience-Yeast Display vs Phage Display

Antibody Discovery,

Yeast display and phage display are two powerful technologies used for protein engineering, antibody discovery, and molecular interaction studies. Both involve displaying proteins, peptides, or antibody fragments on the surface of a biological system (yeast or phage) and screening for high-affinity binders, but they differ in several key aspects. Here’s a comparison of yeast display and phage display:

Biological System

Yeast Display

Uses _Saccharomyces cerevisiae_ (often strain EBY100) or other yeast species.

Proteins or peptides are displayed on the yeast cell surface, typically by fusing them to a cell wall anchor protein, such as Aga2p, which binds to the Aga1p protein on the yeast cell wall.

Yeast cells are eukaryotic, allowing for post-translational modifications (e.g., glycosylation, proper folding) of the displayed proteins.

Phage Display

Uses bacteriophages, usually M13 filamentous phage, as the display system.

Proteins or peptides are fused to one of the phage coat proteins (e.g., pIII or pVIII), which display the proteins on the surface of the phage particle.

Phage display is based on prokaryotic systems (bacteria), meaning that proteins are expressed in Escherichia coli (E. coli) before being displayed on the phage surface. This system does not provide eukaryotic post-translational modifications.

Post-Translational Modifications

Yeast Display

Yeast, being eukaryotic, can perform post-translational modifications like glycosylation, disulfide bond formation, and proper protein folding, which is important for the function and stability of complex eukaryotic proteins (e.g., antibodies, receptors).

Glycosylation in yeast differs from mammalian cells, which can sometimes be a disadvantage if human-like glycosylation is essential for the activity of the displayed protein.

Phage Display

As a bacterial system, phages do not offer eukaryotic post-translational modifications. Proteins that require glycosylation, disulfide bonds, or complex folding may not fold correctly when displayed on phage surfaces.

This limits the ability to display some mammalian proteins and complex molecules.

Screening and Selection

Yeast Display

Selection is typically performed using fluorescence-activated cell sorting (FACS). This allows for quantitative, high-resolution sorting based on the fluorescence intensity of labeled targets, enabling precise selection of yeast cells displaying proteins with high affinity for the target.

Yeast display can be used for iterative rounds of sorting to enrich high-affinity binders, allowing researchers to select for both affinity and expression levels.

Phage Display

Selection is performed using biopanning, where phage particles displaying proteins are exposed to a target (e.g., immobilized on a plate), and those that bind to the target are retained while non-binders are washed away.

Multiple rounds of biopanning help enrich for phages displaying proteins with high affinity for the target, but this method is often less quantitative compared to yeast display with FACS.

Library Size

Yeast Display

Yeast can accommodate libraries of up to 10⁶ to 10⁹ variants, depending on the transformation efficiency. While this is lower than phage display, yeast display still allows for the screening of diverse libraries.

The lower library size is compensated by the high screening precision provided by FACS.

Phage Display

Phage display can handle very large libraries, typically ranging from 10⁹ to 10¹¹ variants or even larger.

The high library size allows for the screening of massive diversity, which is particularly useful for early-stage discovery or when screening highly diverse peptide or antibody fragment libraries.

Ease of Use and Throughput

Yeast Display

Requires specialized equipment like FACS for high-throughput sorting, which provides highly quantitative data but requires access to a flow cytometer.

Yeast culture conditions are relatively straightforward, but expression levels may need optimization, especially for large or complex proteins.

Phage Display

Easier to set up initially as it relies on biopanning, which doesn’t require specialized equipment (just immobilized targets and washing steps).

Higher throughput in terms of the number of variants that can be screened quickly, but lacks the high-resolution screening capability of FACS.

Types of Proteins Displayed

Yeast Display

Works well for displaying large, complex proteins (e.g., full-length antibodies, receptors, enzymes) due to the eukaryotic system’s ability to fold and modify these proteins correctly.

It is particularly useful for antibody discovery, affinity maturation, and screening of protein-protein or protein-peptide interactions.

Phage Display

Commonly used for displaying small peptides, antibody fragments (e.g., single-chain variable fragments, scFv; Fab fragments), or relatively simple proteins.

While phage display can handle small peptides well, complex proteins or full-length antibodies may not fold properly.

Protein Size and Complexity

Yeast Display

Can handle large and complex proteins, including full-length antibodies, extracellular domains of membrane proteins, or multi-domain enzymes, because of its eukaryotic folding and modification capabilities.

Phage Display

Works well for small peptides and simpler proteins, but larger proteins or those requiring post-translational modifications may not fold properly and therefore may not function optimally when displayed on the phage surface.

Applications

Yeast Display

Antibody discovery and engineering: Yeast display is particularly useful for affinity maturation, epitope mapping, and antibody engineering because it allows for high-precision sorting.

Protein engineering: Yeast display is effective in screening libraries of proteins for improved binding, stability, or enzymatic activity.

Receptor-ligand interactions: The ability to display large, complex proteins makes yeast display suitable for studying receptor-ligand interactions

Phage Display

Antibody discovery: Phage display is widely used for the initial discovery of antibody fragments (e.g., scFvs, Fabs) from large libraries due to the system’s ability to handle extremely large libraries.

Peptide display: Phage display is ideal for discovering peptide binders or mimotopes that mimic epitopes recognized by antibodies.

Protein-protein interactions: Phage display is also used for discovering protein-protein interaction domains, especially when screening small, stable protein fragments.

Affinity and Selection Precision

Yeast Display

Provides high precision in affinity selection due to FACS, which can distinguish small differences in binding affinity, making it an excellent choice for fine-tuning protein interactions.

Allows for quantitative analysis of protein binding affinities in real time during the selection process.

Phage Display

Offers good initial screening for binding but lacks the precision of yeast display in distinguishing small affinity differences.

Typically used for broad selection of binders, which may require further optimization in later stages of development.

Speed and Iteration

Yeast Display

Can be slower to develop due to the complexity of yeast culture, expression induction, and FACS-based sorting, especially when working with larger proteins.

However, real-time affinity measurement during FACS sorting can save time in downstream validation.

Phage Display

Faster for initial rounds of selection and screening due to the simple biopanning process, which can be completed quickly.

Iterative biopanning can take several rounds to yield high-affinity binders, followed by downstream validation steps.

 Conclusion: Yeast Display vs. Phage Display

Yeast display is ideal for applications that require high precision and screening of complex or large proteins, making it especially useful for antibody engineering, affinity maturation, and eukaryotic protein interactions. Its strength lies in the ability to perform quantitative screening and handle proteins requiring post-translational modifications.

Phage display excels in high-throughput screening of large libraries, particularly for smaller, less complex proteins or peptides, and is widely used in early-stage antibody discovery and peptide screening. It’s faster for initial screening but may require additional rounds of optimization and validation for complex targets.

The choice between yeast display and phage display depends on the specific application, the size and complexity of the protein being studied, the need for post-translational modifications, and the level of precision required in the selection process.

2011 月/24

KMD Bioscience-Yeast Surface Display Protocol

Antibody Discovery

Yeast surface display (YSD) is a powerful technique for expressing proteins, peptides, or antibody fragments on the surface of yeast cells, allowing researchers to screen and study protein-protein, protein-peptide, or protein-small molecule interactions. The following protocol outlines the steps for constructing and using a yeast surface display system.

 Yeast Surface Display Protocol

 Materials Needed

  1. Yeast strain: _Saccharomyces cerevisiae_ strain (commonly EBY100, which has been optimized for surface display).
  2. Yeast expression vector: Plasmid vector (e.g., pCTCON2 or pYD1) with the gene of interest cloned in frame with the surface display protein (e.g., Aga2p).
  3. Growth media:

SD-CAA (minimal selection media): For plasmid selection.

SG-CAA (induction media): For induction of protein expression (galactose-containing media).

  1. Enzymes and reagents:

DNA extraction kits or methods.

Restriction enzymes for cloning.

  1. Antibiotics: For selection, if applicable.
  2. Fluorescently labeled antibodies/target molecules: Used for detection of displayed proteins.
  3. PBS: Phosphate-buffered saline for washing steps.
  4. Induction buffer: To induce protein expression in yeast.
  5. Plasmid DNA preparation kits.

 Equipment Needed

  1. Shaking incubator: For growing yeast cultures at optimal temperatures.
  2. Centrifuge: For pelleting cells.
  3. Flow cytometer (FACS): For sorting yeast cells based on surface display and target binding.
  4. Microcentrifuge tubes: For handling small volumes of yeast cultures and reagents.
  5. Electroporator or chemical transformation materials: For yeast transformation.

Protocol

Cloning the Gene of Interest into a Yeast Surface Display Vector

Design the gene of interest: The gene encoding the protein or peptide of interest should be cloned into the yeast display vector (e.g., pCTCON2) in frame with the surface anchor protein gene, such as Aga2p.

The vector typically contains a galactose-inducible promoter (GAL1) for controlled expression.

Restriction digestion and ligation: Use appropriate restriction enzymes to cut the vector and the insert (gene of interest), followed by ligation to create the expression construct.

Transformation of E. coli: Transform the ligated plasmid into E. coli for amplification and plasmid preparation. Isolate the plasmid DNA using a DNA preparation kit.

Verification: Sequence the plasmid to verify correct insertion and reading frame with the Aga2p gene.

Transformation of Yeast Cells

Prepare yeast cells: Use _Saccharomyces cerevisiae_ (such as EBY100) as the host strain.

Transform the plasmid into yeast:

Chemical transformation: Use the lithium acetate method or electroporation to transform the plasmid into yeast cells.

Incubate on SD-CAA plates (synthetic defined medium with casein amino acids) that lack uracil or another selectable marker, to select for yeast cells that have taken up the plasmid.

Colony isolation: Grow transformed yeast cells on selective plates at 30°C for 2-3 days until colonies form.

Culture and Induction of Protein Expression

Inoculation and culture:

Inoculate a single colony from the transformation plate into SD-CAA medium and grow overnight at 30°C with shaking (250–300 rpm).

Induction of protein expression:

The next day, pellet the yeast cells by centrifuging at 1,000–3,000 x g for 5 minutes.

Wash cells with sterile water or PBS and resuspend them in SG-CAA medium (containing galactose) to induce the expression of the protein on the yeast surface.

Incubate at 20°C or 30°C for 24–48 hours with shaking to allow for protein display.

Confirmation of Protein Display

Sample preparation: After induction, collect 1 mL of the culture and pellet the yeast cells by centrifugation (1,000–3,000 x g for 5 minutes).

Washing: Wash the cells twice with cold PBS to remove residual media and prevent non-specific binding in downstream assays.

Fluorescent labeling:

Incubate the yeast cells with a primary antibody or fluorescently labeled target molecule that binds to the protein of interest. If using a primary antibody, follow with a secondary fluorescently labeled antibody.

Typically, 30 minutes to 1 hour at 4°C is sufficient for incubation, with gentle mixing to avoid settling of yeast cells.

Washing: After labeling, wash the cells 2-3 times with PBS to remove unbound antibodies or fluorescent probes.

Flow cytometry: Analyze the labeled yeast cells using a flow cytometer (FACS). The presence of fluorescence indicates successful display of the protein on the yeast surface.

Screening for High-Affinity Binders (Optional)

If you are screening for high-affinity binders, such as in antibody discovery or protein engineering:

Incubate yeast cells with target: Incubate the yeast cells with the fluorescently labeled target molecule (e.g., an antigen, protein, or small molecule).

This step helps identify yeast cells displaying proteins or peptides that specifically bind to the target molecule.

FACS sorting: After incubation, use FACS to sort yeast cells that show strong binding to the target (high fluorescence). Collect the sorted cells for further rounds of selection and enrichment.

Iterative rounds: Perform additional rounds of growth, induction, and FACS sorting to progressively enrich for yeast cells displaying proteins or peptides with higher affinity for the target molecule.

Recovery and Sequence Analysis

DNA extraction: After sorting, extract plasmid DNA from the enriched yeast cells to identify the sequences of the selected binders.

Yeast plasmid DNA can be extracted using yeast-specific plasmid prep kits or protocols.

Transformation into E. coli: Transform the extracted plasmid DNA into E. coli for amplification and sequencing.

Sequence analysis: Sequence the plasmids to identify the mutations or variants that led to enhanced binding or functional properties.

Characterization of Selected Variants

Once specific yeast clones have been isolated, express the proteins or peptides in larger quantities for further biochemical and functional characterization:

Binding affinity measurement: Use surface plasmon resonance (SPR) or isothermal titration calorimetry (ITC) to measure the binding affinity of the selected proteins.

Functional assays: Perform any necessary functional assays to validate the biological activity of the selected proteins or peptides.

 Important Considerations

Glycosylation: Yeast cells are eukaryotic, meaning they glycosylate proteins. This is advantageous for many applications, but yeast glycosylation patterns differ from mammalian systems, which can sometimes affect the biological activity of the displayed proteins.

Temperature for induction: Lower temperatures (20°C) may help improve proper protein folding and display efficiency, especially for complex or large proteins.

Antigen/target concentration: Use an optimized concentration of the target molecule to avoid non-specific binding or saturation during sorting or affinity screening.

 Conclusion

Yeast surface display is a powerful tool for protein engineering, antibody discovery, and studying molecular interactions. This protocol provides the steps to construct, express, and screen proteins or peptides on the surface of yeast cells, making it useful for applications ranging from high-affinity binder discovery to functional protein analysis.

1911 月/24

KMD Bioscience-Yeast Display Library

Antibody Discovery

A yeast display library refers to a large collection of diverse protein variants that are displayed on the surface of yeast cells, typically _Saccharomyces cerevisiae_, for screening and selection. These libraries are widely used in protein engineering, antibody discovery, and directed evolution to identify variants with desirable properties, such as increased binding affinity, stability, or specificity.

 Overview of Yeast Display Libraries

Yeast display libraries are constructed by generating a diverse set of genetic sequences encoding the protein or peptide variants of interest. These sequences are then cloned into yeast expression vectors that fuse the protein with a surface anchoring protein, such as Aga2p, allowing the proteins to be displayed on the yeast cell surface. The diversity of the library can range from thousands to billions of variants.

Once the library is constructed, it can be screened against specific targets, like antigens or ligands, using techniques like fluorescence-activated cell sorting (FACS) to isolate high-affinity binders or proteins with specific characteristics.

 Steps in the Construction and Use of a Yeast Display Library

Library Design and Generation

The first step in creating a yeast display library is generating diversity at the genetic level. This diversity can be introduced through several methods:

Random Mutagenesis: Mutations are introduced into the gene of interest through error-prone PCR or chemical mutagenesis, creating random variations in the sequence.

Recombination-Based Libraries: Combining fragments from related genes or domains to create hybrid proteins. This is often used in antibody or enzyme engineering to mix and match different functional domains.

Synthetic Libraries: Custom-designed sequences are synthesized to introduce specific amino acid changes, either at key residues or across an entire protein.

Natural Repertoires: Libraries can also be created from natural sources, such as B-cell repertoires in antibody discovery, where variable regions of antibody genes are amplified and cloned into the yeast display system.

Cloning into Expression Vectors

Once the genetic diversity is created, the variants are cloned into an expression vector. These vectors typically contain:

Promoter: To drive protein expression (e.g., galactose-inducible promoter).

Protein of Interest: The gene encoding the protein variants is fused to a surface anchor protein (e.g., Aga2p) so that the protein is displayed on the yeast cell wall.

Epitope Tags: Optional tags like HA or FLAG can be added to the protein for easy detection and quantification during screening.

Selection Marker: Antibiotic resistance or other selection markers are included to ensure that only yeast cells containing the plasmid grow during the selection process.

Transformation of Yeast Cells

The library of plasmids is then introduced into yeast cells by high-efficiency transformation methods such as electroporation or chemical transformation. The goal is to generate a yeast population where each cell expresses a different protein variant on its surface. The size of the library is determined by how many unique clones are transformed into the yeast cells, typically ranging from 10⁶ to 10¹¹.

Induction of Protein Display

Once transformed, the yeast cells are grown under conditions that induce the expression of the gene of interest and its display on the cell surface. Common induction systems use galactose-inducible promoters to ensure high levels of surface display.

The protein is fused to a yeast surface protein (Aga2p), which is attached to the cell wall by binding to Aga1p, a component of the yeast cell wall machinery.

Screening the Yeast Display Library

Once the library is constructed and the proteins are displayed on the surface of the yeast cells, the next step is to screen for variants that have the desired properties. The most common technique used is fluorescence-activated cell sorting (FACS), which allows high-throughput sorting based on binding affinity to a target antigen or ligand.

Binding to Target: The yeast cells are incubated with a fluorescently labeled target molecule (e.g., an antigen or ligand). Cells displaying proteins that bind to the target will have the fluorescent signal on their surface.

FACS: Fluorescent cells are sorted by FACS, allowing researchers to isolate those yeast cells that display protein variants with high affinity or specificity for the target.

Library Enrichment: After each round of sorting, the selected yeast cells are cultured, and the process can be repeated over several rounds to progressively enrich for the best-performing variants.

Recovery and Sequencing of High-Affinity Variants

Once high-affinity binders or variants with desired properties are selected, the yeast cells can be lysed, and the plasmid DNA encoding the protein variants can be extracted. The recovered DNA is sequenced to identify the mutations or variations in the protein sequence that led to improved binding or other traits.

Next-Generation Sequencing (NGS): NGS can be used to comprehensively analyze the selected clones and assess the diversity and frequency of specific variants.

Validation of Selected Variants

After screening and sequencing, the selected protein variants are typically validated in further biochemical or functional assays. These might include:

Binding Affinity Measurements: Techniques such as surface plasmon resonance (SPR) or isothermal titration calorimetry (ITC) can be used to precisely measure the binding affinity of the selected proteins to their target.

Functional Assays: For enzymes or receptors, functional assays can confirm that the variant performs its intended biological function (e.g., catalysis or signaling).

Protein Expression and Purification: The selected variants can be expressed and purified in larger quantities to study their properties in vitro or in vivo.

 Applications of Yeast Display Libraries

Antibody Discovery and Engineering

Yeast display libraries are extensively used to screen large combinatorial libraries of antibody fragments (such as scFvs or Fabs) to identify high-affinity binders. This is a key technique in antibody development for therapeutic and diagnostic purposes.

Protein Engineering and Directed Evolution

By screening libraries of enzyme variants, researchers can evolve enzymes to have improved catalytic efficiency, altered substrate specificity, or enhanced stability under different conditions.

Peptide Library Screening

Libraries of short peptides can be displayed on yeast to identify binding motifs or develop peptide-based therapeutics.

Study of Protein-Protein Interactions

Yeast display allows researchers to screen for and study protein-protein interactions, helping to map interaction domains or design proteins with enhanced interaction capabilities.

Receptor-Ligand Screening

Yeast display can be used to display receptors, and libraries can be screened to identify ligands or to study the interaction between receptors and their binding partners.

 Advantages of Yeast Display Libraries

Eukaryotic System: Yeast cells are eukaryotic, which means they can carry out post-translational modifications and proper folding of displayed proteins, unlike bacterial systems (e.g., phage display).

High-Throughput: Yeast display libraries can be screened in high-throughput using FACS, allowing rapid identification of the best variants.

Quantitative Screening: FACS allows for quantitative assessment of binding affinity, making it possible to rank protein variants based on their performance.

Library Size: Yeast display can handle large libraries, up to 10⁸-10⁹ variants, providing significant diversity for selection.

 Limitations of Yeast Display Libraries

Protein Size: Large or complex proteins may not display efficiently on the yeast surface due to size or folding constraints.

Glycosylation Differences: Yeast glycosylation patterns differ from those in humans, which may affect the functionality of glycosylated proteins displayed on the yeast surface.

Transformation Efficiency: Although yeast can handle large libraries, transformation efficiency can sometimes be a bottleneck compared to other systems like phage display.

 Conclusion

Yeast display libraries are a powerful tool for protein engineering, antibody discovery, and studying molecular interactions. By screening vast libraries of protein variants on the surface of yeast cells, researchers can rapidly identify and optimize proteins with desired properties for therapeutic, diagnostic, or industrial applications.

1811 月/24

KMD Bioscience-Yeast Surface Display

Antibody Discovery

Yeast surface display (YSD) is a molecular technique used to display proteins, peptides, or antibody fragments on the surface of yeast cells. This method allows researchers to study the binding properties of proteins, perform directed evolution for improving affinity or specificity, and screen for protein-protein interactions. Yeast surface display is particularly useful in antibody engineering and protein evolution due to the ease of screening large libraries of variants in a high-throughput manner.

 How Yeast Surface Display Works

Expression of Proteins on the Yeast Surface

The process begins by expressing a protein or peptide of interest on the surface of the yeast cells (commonly _Saccharomyces cerevisiae_). The gene encoding the protein is fused to genes that encode surface proteins, such as Aga2p, which is naturally located on the yeast cell wall. This allows the protein of interest to be anchored on the outer surface of the yeast cell for interaction studies.

Fusion to Aga2p: The protein of interest is genetically fused to the Aga2p protein, which then binds to the Aga1p subunit present on the yeast cell wall. This anchoring mechanism allows the protein to be displayed externally, enabling direct interactions with target molecules.

Display of Libraries: Libraries of protein variants can be generated and displayed on the surface of yeast cells. These variants may differ by just a few amino acids, making it possible to study the effect of mutations on binding affinity or specificity.

Screening and Selection

Once the proteins are displayed on the yeast surface, their interactions with specific ligands or other proteins can be assessed. The process usually involves:

Fluorescence-Activated Cell Sorting (FACS): Yeast cells displaying proteins are incubated with fluorescently labeled ligands, antigens, or antibodies. The cells that bind to the fluorescent ligands can be sorted based on fluorescence intensity, allowing for the selection of yeast cells that display proteins with high binding affinity.

Library Screening: Large libraries of proteins or antibody fragments can be screened using FACS to identify variants that bind to a target molecule with improved properties (e.g., higher affinity or specificity). Multiple rounds of sorting and mutagenesis can be used to perform directed evolution, a process that iteratively improves the desired properties of the protein.

Analysis and Validation

After sorting, the selected yeast cells can be further analyzed to identify the best variants:

DNA Sequencing: The genes encoding the selected proteins are sequenced to determine the mutations that contributed to improved binding properties.

Binding Assays: Surface-displayed proteins are subjected to detailed binding assays (such as affinity measurements) to validate their performance. These assays may involve quantitative fluorescence measurements, competition binding experiments, or flow cytometry.

 Applications of Yeast Surface Display

Antibody Engineering

Affinity Maturation: YSD is widely used for affinity maturation of antibody fragments, such as scFv (single-chain variable fragment) and Fab (fragment antigen-binding). Libraries of antibody variants are displayed on yeast, and FACS is used to isolate clones with enhanced binding to antigens.

Screening Antibody Libraries: Yeast surface display allows for the selection of antibodies from large combinatorial libraries based on their binding specificity to target antigens.

Protein-Protein Interactions

YSD can be used to identify and study protein-protein interactions. By displaying one protein on the yeast surface and screening for its interaction with other proteins or ligands, researchers can map interaction sites or design proteins with enhanced binding affinity.

Enzyme Engineering

Directed evolution can be applied to enzymes displayed on yeast cells to improve properties such as catalytic efficiency, stability, or substrate specificity.

Therapeutic Protein Development

YSD is used to engineer and optimize therapeutic proteins like cytokines, growth factors, and other biologics by screening for variants with improved stability, reduced immunogenicity, or enhanced activity.

Receptor-Ligand Studies

The technique allows for the study of receptor-ligand interactions, which is important for understanding signaling pathways or developing drugs that target specific receptors.

Vaccine Development

Yeast surface display has been used in the development of vaccines by displaying antigens on yeast cells, which can help elicit immune responses when used in immunization strategies.

 Advantages of Yeast Surface Display

High-Throughput Screening: YSD allows for the rapid screening of large libraries of protein variants, enabling the selection of optimal candidates in a relatively short amount of time.

Quantitative Analysis: FACS-based screening provides quantitative data on binding affinities, making it possible to accurately compare different variants.

Post-Translational Modifications: Yeast cells are eukaryotic, allowing for proper folding and post-translational modifications (such as glycosylation) of the displayed proteins, making them more biologically relevant compared to bacterial systems.

Non-Toxic Display: Since the protein is displayed on the surface, yeast surface display avoids potential issues with toxicity that can arise when proteins are expressed internally in cells.

 Limitations of Yeast Surface Display

Size Limitation: Larger proteins may be challenging to display due to limitations in yeast secretion pathways and the ability of the protein to fold correctly on the surface.

Glycosylation Differences: Yeast glycosylation patterns can differ from those in higher eukaryotes like mammals, which may affect the function of glycosylated proteins.

Library Size: While YSD can handle large libraries, the overall diversity may still be lower than in other systems like phage display due to the limitations of transformation efficiency in yeast.

Conclusion

Yeast surface display is a versatile and powerful tool for protein engineering, antibody discovery, and studying molecular interactions. Its ability to screen large libraries for high-affinity variants and its use in directed evolution make it an invaluable method in research and biotechnology for developing proteins with desired properties.

1211 月/24

KMD Bioscience-Primary Cell Culture Protocol

CRO Service

Primary cell culture involves the isolation and growth of cells directly from tissues cultured in vitro for experimental research. Primary cells reflect the biological characteristics of the tissue they are derived from, making them a valuable tool in biological research, drug testing, and disease modeling. Below is a detailed protocol for setting up primary cell cultures.

Primary Cell Culture Protocol

 Materials Needed

1. Tissue Source: Organ or tissue from which cells are to be isolated (e.g., liver, lung, skin, etc.).

2. Enzymes for Tissue Dissociation

Collagenase (typically used for soft tissues).

Trypsin (to detach cells from tissues or culture plates).

DNase (to break down DNA released from dead cells).

Dispase or hyaluronidase (for tougher tissues).

3. Culture Media

Choose a culture medium appropriate for the cell type (e.g., DMEM, RPMI-1640, or specialized media such as Keratinocyte Serum-Free Media (KSFM) or Endothelial Cell Growth Media).

Supplements like FBS (Fetal Bovine Serum), growth factors (EGF, bFGF), antibiotics (penicillin-streptomycin), and glutamine.

  1. Cell Strainers (40-70 µm) to filter cell suspension.
  2. Sterile Petri Dishes, culture plates, or flasks (depending on the intended culture format).
  3. Sterile PBS (phosphate-buffered saline).
  4. Trypsin-EDTA or enzyme-free dissociation buffer (if necessary).
  5. Centrifuge.
  6. Laminar Flow Hood and Incubator: Standard tissue culture equipment to maintain sterility and temperature (typically 37°C with 5% CO₂ for mammalian cells).

 Procedure

Preparation and Sterilization

Sterilize all tools: Dissecting instruments (scalpels, scissors, forceps), Petri dishes, and pipettes should be sterilized by autoclaving or using ethanol.

Work in a sterile environment: All procedures should be conducted under a laminar flow hood to maintain sterility.

Prepare the culture medium: Warm the appropriate medium supplemented with FBS, antibiotics, and any necessary growth factors to 37°C in advance.

Tissue Collection and Dissection

Collect the tissue: Immediately after excision, place the tissue in a sterile container with cold PBS or transport media (such as HBSS or DMEM) to keep it viable during transfer.

If obtaining tissue from an animal, ensure proper ethical approvals and protocols for handling the tissue.

Dissect the tissue: Under sterile conditions in the hood, cut the tissue into small pieces (~1–2 mm³) using sterile scalpels or scissors. The finer the pieces, the more efficient the digestion process will be.

Tissue Digestion and Cell Isolation

Enzymatic Digestion: To dissociate cells from the tissue matrix:

Transfer the minced tissue into a sterile conical tube.

Add digestion enzymes (e.g., collagenase, trypsin) prepared in warm PBS or culture medium.

Example: For collagenase digestion, use 1-2 mg/mL of collagenase in PBS.

Incubate the tissue with the enzyme solution at 37°C in a shaking water bath or incubator for 30 minutes to 2 hours, depending on the tissue type.

Agitate the tube occasionally to promote dissociation.

After incubation, gently pipette the solution up and down to help further dissociate the tissue into single cells.

Filtration and Centrifugation

Filtration: Pass the cell suspension through a sterile cell strainer (40–70 µm) into a new sterile tube to remove undigested tissue fragments.

Centrifugation: Centrifuge the filtered cell suspension at 300–400 x g for 5-10 minutes at room temperature.

This step will pellet the cells, allowing you to discard the supernatant containing the digestion enzymes and other debris.

Resuspend the Pellet: After centrifugation, carefully aspirate the supernatant and resuspend the cell pellet in fresh culture medium.

Seeding and Culturing Cells

Cell Counting (Optional): At this point, count the cells using a hemocytometer and trypan blue staining to assess cell viability and ensure accurate seeding density.

Seeding: Seed the cells into sterile culture flasks, dishes, or plates at an appropriate density (e.g., 5 x 10⁴ to 1 x 10⁶ cells depending on the cell type and experiment).

Use specialized-coated plates (e.g., collagen, fibronectin, or gelatin-coated plates) for certain cell types, such as endothelial or epithelial cells, which require adherence support.

Incubation: Place the culture flasks or plates in a 37°C incubator with 5% CO₂. Ensure proper culture conditions specific to the cell type being used (e.g., low oxygen for certain primary cell types).

Monitoring Cell Growth

Observe Cells: Over the next 24-48 hours, check the cells regularly under a microscope to monitor attachment, spreading, and growth.

Primary cells typically grow slower than immortalized cell lines, so they may take a few days to adhere and proliferate.

Medium Change: Replace the culture medium every 2-3 days, carefully aspirating the old medium and replacing it with fresh, pre-warmed medium.

Ensure the cells remain in a healthy monolayer and are not over-confluent (typically 70-90% confluency is ideal for subculturing).

Subculturing (Passaging)

Subculturing: When the cells reach appropriate confluence (~70-90%), they should be passaged to avoid overgrowth and maintain their health.

Detachment: Gently wash the cells with sterile PBS to remove any residual medium.

Add trypsin-EDTA or a gentle dissociation buffer to detach the cells. Incubate at 37°C for 2-5 minutes, monitoring under the microscope until the cells round up and begin detaching.

Neutralization: Neutralize the trypsin with fresh complete medium containing serum (FBS). Pipette up and down to create a single-cell suspension.

Centrifuge the cell suspension at 300–400 x g for 5 minutes.

Reseeding: Resuspend the cells in fresh medium and reseed them into new culture dishes at a lower density (e.g., 1:3 or 1:5 split ratios, depending on the cell type).

Cryopreservation (Optional)

If you wish to store primary cells for future use, you can cryopreserve them:

Freezing Medium: Prepare freezing medium (10% DMSO in FBS or complete culture medium).

Cryovials: Aliquot 1–2 million cells per cryovial in freezing medium.

Freezing: Gradually freeze the cells by placing them in a controlled-rate freezing container (-80°C freezer) overnight before transferring them to liquid nitrogen for long-term storage.

 Critical Tips and Considerations

Tissue-Specific Conditions: Each tissue type has specific requirements for dissociation enzymes, media, and growth conditions. Consult protocols or literature specific to your cell type of interest.

Sterility: Maintaining sterility throughout the procedure is critical to avoid contamination.

Cell Type-Specific Media: Primary cells often require specialized growth media with specific supplements like growth factors (e.g., EGF, insulin, hydrocortisone).

Lifespan: Primary cells have a limited lifespan in culture, meaning they can only be passaged a finite number of times before undergoing senescence. Avoid too many passages to maintain the biological relevance of the cells.

 Conclusion

Primary cell culture is a valuable tool for studying physiological processes in a more relevant context compared to immortalized cell lines. Following the steps in this protocol will allow for the successful isolation, culture, and maintenance of primary cells, although specific modifications may be needed depending on the tissue type and cell characteristics. Proper handling, growth conditions, and regular monitoring are essential for preserving the health and viability of primary cells.

0811 月/24

KMD Bioscience-CRISPR-cas9 Knockout Protocol

CRO Service

CRISPR-Cas9 knockout protocols are designed to generate precise gene knockouts by introducing targeted mutations through double-stranded DNA breaks, followed by error-prone repair mechanisms like non-homologous end joining (NHEJ). Below is a typical step-by-step CRISPR-Cas9 knockout protocol, covering the design, delivery, and validation stages.

 CRISPR-Cas9 Knockout Protocol

 Materials

  1. Cell line: Choose the cell line you want to modify (e.g., HEK293T, CHO, etc.).
  2. Plasmids:

Cas9 expression vector (or a cell line that stably expresses Cas9).

gRNA expression vector (e.g., in plasmid form).

Optional: Repair template if using homology-directed repair (HDR) for precise modifications.

  1. Lipofection reagent: For transfection (e.g., Lipofectamine, PEI).
  2. Selection antibiotic: If using selection markers (e.g., puromycin, G418).
  3. Primers: For PCR validation of gene editing.
  4. Next-generation sequencing (NGS) services or sequencing tools: For validation of mutations.
  5. qPCR reagents: For expression analysis if needed.
  6. Antibodies: For protein expression validation, if necessary.

Procedure

Design and Selection of gRNA

Design gRNAs: Design guide RNAs (gRNAs) that specifically target the gene of interest. Most protocols use online tools like:

CRISPR design tools from the Broad Institute (https://portals.broadinstitute.org/gpp/public/analysis-tools/sgrna-design)

Benchling or CHOPCHOP

Target Exon(s): Choose gRNAs targeting an essential exon in the gene to ensure a knockout. The gRNA should direct Cas9 to create a double-strand break (DSB).

Target sequences should have a 5’-NGG-3’ PAM (protospacer adjacent motif) sequence.

Off-Target Analysis: Ensure minimal off-target effects by selecting gRNAs with high specificity using design tools.

Cloning of gRNA into Expression Vector

Clone the selected gRNA sequence into a plasmid vector expressing both the gRNA and Cas9, or into separate plasmids.

Vectors like pSpCas9(BB)-2A-Puro (PX459) from Addgene are commonly used.

Optional: Use a plasmid with a fluorescent marker (e.g., GFP) for transfection efficiency tracking.

Transfection of Cells

Seed Cells: Seed the cells in a 6-well plate or another appropriate format so that they are 50-70% confluent at the time of transfection.

Transfection: Transfect the cells with the gRNA-Cas9 plasmid (and optionally, a donor template for HDR) using a lipofection reagent (e.g., Lipofectamine or PEI) or electroporation.

Amount of DNA: Use 1-2 µg of plasmid DNA per well of a 6-well plate.

Incubation: Incubate the cells for 24-48 hours to allow for gRNA expression and Cas9-mediated DNA cleavage.

Selection of Transfected Cells (Optional)

Antibiotic Selection: If your plasmid contains an antibiotic resistance gene (e.g., puromycin or G418 resistance), apply the appropriate antibiotic to select for successfully transfected cells.

For puromycin selection, use concentrations between 1-10 µg/mL based on the cell type and perform selection for 2-3 days until only transfected cells survive.

 Single-Cell Cloning

Dilution or FACS Sorting: Once the cells are transfected and selected, dilute the cells into 96-well plates to obtain single-cell clones. Alternatively, use FACS to sort GFP-positive cells (if using a fluorescent marker).

Seed the cells at approximately 1 cell per well in a 96-well plate for single-cell isolation.

Allow the clones to grow for 1-2 weeks, replacing the media as needed.

Validation of Knockout

Genomic DNA Extraction

Extract genomic DNA from the single-cell clones using a DNA extraction kit (or traditional phenol-chloroform methods).

PCR and Sanger Sequencing

PCR Amplification: Use gene-specific primers flanking the target site to amplify the region where the Cas9 cut was made.

Run PCR products on a gel to check for indels (insertions or deletions) by size differences.

Send the PCR products for Sanger sequencing to confirm the presence of indels, which are indicative of successful gene knockout.

  T7E1 Assay (Optional)

The T7 Endonuclease I assay detects mismatches in DNA, which are indicative of indels from CRISPR editing. This is done by hybridizing and re-annealing wild-type and mutant sequences, which are then cleaved by T7 Endonuclease I.

Run the digested products on a gel to confirm cleavage, which suggests gene editing.

Western Blot (for Protein Knockout)

To confirm the absence of the target protein, perform a Western blot using specific antibodies against the protein of interest. A reduction or complete absence of the protein suggests a successful knockout at the functional level.

qPCR (for mRNA Knockout)

To confirm loss of gene expression at the transcript level, extract RNA and perform reverse transcription followed by qPCR. The absence or significant reduction in mRNA expression is another indication of successful gene knockout.

Expansion and Further Analysis

Once knockout clones are validated, expand the positive clones for further experiments.

Use flow cytometry, functional assays, or any other downstream analysis depending on the purpose of your knockout study.

 Troubleshooting

  1. Low Transfection Efficiency: Optimize transfection conditions, such as adjusting DNA amounts, using different transfection reagents, or increasing cell density.
  2. No Detected Indels: Ensure that gRNAs target the correct location and that the PAM sequence is correct. Redesign the gRNAs if necessary. Check Cas9 expression using Western blot or qPCR.
  3. Off-Target Effects: Design gRNAs with minimal predicted off-target effects. Use more specific Cas9 variants, like Cas9-HF (high fidelity), if needed.
  4. Cell Death: Knockout of an essential gene might cause cell death. Perform short-term experiments (before cells die) or use conditional Cas9 systems or inducible gRNAs.

 Conclusion

The CRISPR-Cas9 knockout protocol is a powerful tool to generate gene knockouts in a wide range of cell lines. By following the outlined steps—starting with gRNA design, transfection, and selection, followed by validation through PCR, sequencing, and protein assays—you can efficiently create knockout cell lines for research. Proper validation of gene editing, including confirmation of off-target effects and functional loss, is crucial for the success of these experiments.