Experimental methods summary (2): cell biology
1. Primary cell isolation and culture ... 2
1.1. Primary cell isolation ... 2
1.2. Primary cell culture ... 2
2. Stem-cell isolation and culture ... 3
2.1. Directed differentiation of embryonic stem cells ... 3
2.2. Cancer stem-cell (CSC) isolation and identification ... 3
3. Cell-proliferation methods ... 5
3.1. MTT assay ... 5
3.2. XTT assay ... 5
3.3. WST-1 assay ... 6
3.4. CCK-8 (WST-8) assay ... 6
3.4. Colony formation in plates ... 6
3.5. Soft-agar colony formation ... 7
3.6. BrdU assay ... 7
4. Cell migration and invasion methods ... 8
4.1. Scratch assay ... 8
4.2. Cell adhesion ... 8
4.3. Transwell migration ... 9
4.4. Transwell invasion ... 9
5. Angiogenesis ... 10
6. Cell cycle: PI staining ... 10
7. Apoptosis assays ... 10
7.1. Annexin V/PI double staining ... 10
7.2. TUNEL assay ... 11
7.3. Mitochondrial membrane potential (MMP) ... 12
8. Cell sorting ... 12
8.1. Flow sorting ... 12
8.2. Magnetic-bead sorting ... 13
9. Cell transfection ... 13
10. Lentiviral infection of cells ... 14
11. Selection of stable cell lines ... 14
12. Selection of drug-resistant cell lines ... 15
13. Cell co-culture ... 15
13.1. Direct co-culture ... 15
13.2. Indirect co-culture ... 15
Tissues from humans or animals, including embryonic tissue, contain tightly associated cells. To culture cells individually in vitro, disperse the tissue and dissociate the cells.
First: isolation of cells in suspension.
For suspensions such as blood, amniotic fluid, pleural fluid or ascites, the source describes low-speed centrifugation at 1,000 r/min for 10 min. Different cell densities may form layers in a separation medium, from which the desired cells can be collected.
Second: isolation from solid tissue.
Solid tissues can be dissociated mechanically or by digestion. Mechanical dispersion is simple and rapid but may damage tissue and disperse cells poorly; the source describes it for soft tissue with little fibrous material. Digestion cuts tissue into small pieces and uses enzymatic or nonenzymatic treatment to loosen intercellular connections. The loosened material can then be dispersed mechanically by pipetting, stirring or agitation to produce a suspension of small cell clusters and individual cells for culture.
Primary culture is the first in vitro culture of cells taken from donor tissue and the first step in establishing a cell line.
First: tissue-explant culture.
In explant culture, small pieces of tissue are placed in a culture flask or dish. A thin collagen coating may be applied to help the tissue adhere so cells can grow outward along the surface.
Second: culture after tissue digestion.
Third: suspension-cell culture.
Suspension cells, such as leukemia cells, lymphocytes, bone-marrow cells, malignant cells in pleural fluid or ascites, and immune cells, can be separated by low-speed centrifugation and cultured directly or separated on lymphocyte separation medium before plating.
Fourth: organ culture.
Organ culture places an organ or tissue piece in a defined in vitro environment without first dissociating it. It preserves relative tissue organization for examining cell arrangement, interactions and local regulatory effects.
The source describes dissociating embryoid bodies into single cells, allowing attachment and applying stage-dependent chemicals, conditioned media or cytokines to promote or inhibit differentiation. It describes three approaches: adding growth factors or chemical inducers, co-culturing embryonic stem (ES) cells with other cells, and culturing on suitable substrates to affect gene expression and lineage differentiation.
The source groups in vitro ES-cell induction into chemical, cytokine and exogenous-gene approaches.
Chemical inducers listed include retinoic acid (RA), DMSO, β-glycerophosphate, vitamin C, dexamethasone, vitamin K3 and “2,5-hydroxyvitamin D3.” The last name is reproduced from the source and requires verification.
Cytokine conditions can influence ES-cell proliferation or differentiation. The source identifies bone morphogenetic proteins (BMPs) and fibroblast growth factors (FGF) among studied factors.
Exogenous-gene approaches use expression of a differentiation-promoting gene to influence ES-cell fate. The source emphasizes identifying relevant genes and ensuring appropriate timing and genomic insertion.
The historical source groups cancer stem-cell (CSC) isolation approaches by cell phenotype and biological properties.
Category 1: isolation based on cell-surface markers
Hematologic cancer stem cells: the source lists the normal hematopoietic stem-cell phenotype as CD34+CD38+Thy-1−. This historical description requires verification and should not be treated as a definitive marker panel.
Cancer stem cells from solid tumors:
A. Breast cancer: the source describes ESA+CD44+CD24−/low Lineage− cells as breast CSCs; marker-based identification depends on the study and requires validation.
B. Brain tumors: the source describes CD133+ tumor cells as brain tumor stem cells (BTSCs) and lists Sox2, Musashi-1, bmi-1 and a phosphoserine phosphatase among associated neural/stem-cell features. These historical marker claims require validation.
C. Colorectal cancer: the source describes CD133+ cells as tumor-initiating cells and lists epithelial cell adhesion molecule, CD44 and CD166 as potential surface markers; applicability requires validation.
Example: breast cancer (adapt the blue terms for other tumor types):
Isolation and culture of human breast tumor cells: with informed consent from the patient and family, obtain about 10 g of tumor tissue under sterile operating-room conditions from a breast cancer patient at XXX hospital. Transfer promptly in sterile transport solution, wash several times with PBS, remove fat and necrotic tissue, and mince. Add 8 mL DMEM digestion mixture containing 1 mL type IV collagenase at 2,700 U/mL, 100 μL hyaluronidase at 0.02 mg/mL and 50 μL DNase; mix gently and digest at 37°C for 1–2 h, shaking every 20–30 min. Add 10 mL PBS, centrifuge at 1,000 r/min for 4 min and discard the supernatant. Add 1 mL breast tumor stem-cell medium (DMEM-F12 with 10 mL/L FBS, 100 U/mL penicillin, 100 mg/mL streptomycin, 10 ng/mL bFGF, 20 ng/mL EGF and 20 ng/mL ITS), mix, centrifuge at 1,000 r/min for 4 min and discard the supernatant. Seed at 1×10⁶/mL and culture at 37°C with 50 mL/L CO2. Change medium after 3 days; suspension spheres reportedly appear in about 7–10 days. The streptomycin concentration and ITS unit are reproduced as written and require verification.
Flow-cytometric detection of breast tumor cells: collect cultured breast tumor spheres, centrifuge and discard medium. Wash once with PBS, digest with Accutase for 2 min and stop with an equal volume of medium. Transfer to a 15 mL tube, centrifuge at 1,000 r/min for 4 min, resuspend in PBS and filter through a sterile membrane. In four tubes, add 10 μL of each labeled-antibody combination: IgG1-PE/IgG2-FITC, CD44-FITC/IgG1-PE, CD24-PE/IgG2-FITC and CD44-FITC/CD24-PE. Add 500 μL cell suspension to each tube, mix, incubate at room temperature in the dark for 15 min and analyze by flow cytometry.
Second category: side-population (SP) isolation based on efflux of the DNA-binding dye Hoechst 33342.
The source describes a weakly stained subset of Hoechst 33342-treated bone-marrow cells that appears to one side of the main hematopoietic stem/progenitor-cell population on a two-color plot after UV excitation. It reports blue and red detection at 450 nm and 675 nm, respectively, and fewer than 0.1% of cells in the side population. These are historical source statements.
Experimental method:
(1) Staining: prepare a single-cell suspension in DF12 prewarmed to 37°C at 1×10⁶/mL. Add Hoechst 33342 to 6 μg/mL, incubate at 37°C in the dark for 90 min, and place on ice for 10 min. Count, centrifuge, wash and resuspend in ice-cold DF12; adjust sorting samples above 1×10⁷/mL and add 1× DNase I if needed. Before analysis add PI to 2 μg/mL and filter through the “400” filter stated by the source; its unit is unspecified and must be checked. (2) Flow cytometry: the source specifies 355 nm UV excitation, a 610 nm short-pass dichroic and 450/675 nm filters. Evaluate forward and side scatter, plot Hoechst Red versus Hoechst Blue and define an SP gate as the low-blue/low-red region absent in the verapamil control, then calculate and sort SP and non-SP cells.
The historical source describes MTT as a yellow tetrazolium dye reduced by viable cells to blue formazan crystals, with the amount used as an indirect measure of viable cell number. The stated mechanism and proportionality depend on assay conditions.
Experimental method:
Prepare a single-cell suspension of XXX cells in medium containing 10% fetal calf serum. Seed 1,000–10,000 cells per well in a 96-well plate at 200 μL per well and culture for 3–5 days as appropriate. Add 20 μL of 5 mg/mL MTT in PBS per well and incubate for 4 h. Remove supernatant carefully; centrifuge suspension cells first. Add 150 μL DMSO per well, shake for 10 min to dissolve crystals, and measure absorbance at 490 nm. Plot absorbance against time.
The source describes XTT as a tetrazolium derivative reduced by viable cells to a water-soluble brown formazan product; with an electron-coupling reagent, product formation is used as an indirect measure of proliferation.
Experimental method:
The source specifies 6.6 mmol/L XTT and 220 mmol/L phenazine methosulfate (PMS), mixed 1:1 immediately before use. Prepare XXX cells in medium with 10% fetal calf serum and seed 1,000–10,000 cells per well at 200 μL in a 96-well plate. Culture 3–5 days, add 20 μL XTT/PMS per well 2 h before the endpoint, mix, incubate 2 h and measure at 450 nm with a 655 nm reference. The unusually stated stock concentrations are reproduced as written and require verification against the actual reagent protocol.
The source describes WST-1 as a nonradioactive colorimetric assay for viable cells in proliferation or cytotoxicity studies, and claims greater sensitivity and a wider linear range than several other tetrazolium assays; those comparative claims depend on conditions.
Experimental method:
Seed approximately 1×10⁴ XXX cells in 100 μL per well in a 96-well plate. Culture at 37°C and 5% CO2 for 24 h, add test compound at the chosen concentration, and incubate for the desired period at 37°C, 5% CO2 and stated 100% humidity. Add 10 μL WST-1 working solution per well, incubate 1–4 h at 37°C and read absorbance at 450 nm.
The source describes CCK-8 as a water-soluble tetrazolium assay similar to WST-1, producing an orange-yellow formazan product in the presence of an electron carrier. Absorbance is used as an indirect measure of viable-cell number within the assay’s validated range.
Experimental method:
Seed 100 μL cell suspension per well in a 96-well plate and preincubate at 37°C and 5% CO2. After treatment, add 10 μL CCK solution per well, avoiding bubbles that affect optical-density readings. Incubate 1–4 h and read at 450 nm. The source states that, if reading is delayed, 10 μL of 0.1 M HCl or 1% w/v SDS may be added per well and the covered plate kept at room temperature in the dark for measurement within 24 h; verify against the kit instructions.
The source describes plate colony formation as suitable for adherent cells and emphasizes a well-dispersed single-cell suspension and appropriate seeding density.
Experimental method:
Digest logarithmically growing monolayer XXX cells with 0.25% trypsin, disperse and suspend in RPMI 1640 with 10% fetal bovine serum. Dilute and seed, for example, 50, 100 or 200 cells per dish in 10 mL medium prewarmed to 37°C. Distribute evenly and culture at 37°C, 5% CO2 and saturated humidity for 2–3 weeks. Once colonies are visible, remove supernatant, wash twice with PBS and fix for 15 min in 5 mL methanol or 1:3 acetic acid/methanol. Stain with Giemsa for 10–30 min, wash slowly and air dry. Count colonies by eye on a grid or microscopically when they contain more than 50 cells. Colony formation rate = colonies / cells seeded × 100%.
The source describes soft-agar colony formation for tumor or transformed cell lines, states agar should not exceed 40°C when mixed with cells, and gives a maximum of 35 cells/cm² and an example of 1,000 cells per 6 cm dish. It states that normal cells do not proliferate in suspension; this generalization should not be treated as universal.
Experimental method:
Digest logarithmically growing XXX cells with 0.25% trypsin, disperse to single cells, count viable cells and adjust in DMEM with 20% fetal bovine serum. The source states 1×10⁶ cells/L; verify this unit and intended concentration before use. Prepare 1.2% and 0.7% low-melting agarose, sterilize and keep at 40°C. Mix 1.2% agarose 1:1 with 2× DMEM containing 2× antibiotics and 20% calf serum; add 3 mL to a 6 cm dish (7–10 mL to a 10 cm dish), allow to solidify and hold in a CO2 incubator. Mix 0.7% agarose 1:1 with 2× DMEM, add 0.2 mL cell suspension and overlay the bottom layer. After solidification, culture at 37°C and 5% CO2 for 10–14 days and count colonies with an inverted microscope. The source later refers to the bottom layer as 1.2% despite the stated 1:1 dilution; verify the final agar concentration.
BrdU is described as a thymine analog incorporated into newly synthesized DNA in proliferating cells during S phase. Anti-BrdU antibodies can detect incorporation as an indicator of proliferation.
Experimental method:
Seed XXX cells at 1.5×10⁵/mL in a 35 mm dish containing a coverslip. Culture for one day, then synchronize for 3 days in 0.4% FBS medium to place most cells in G0. Add BrdU (1.0 mg/mL stock, 0.03 μg/mL final), incubate at 37°C for 40 min, discard medium and wash three times with PBS. Fix with methanol/acetic acid for 10 min, air-dry, treat with 0.3% H2O2 in methanol for 30 min and block with 5% normal rabbit serum. Denature nucleic acids with formamide at 100°C for 5 min, cool on ice and wash with PBS. Add anti-mouse BrdU monoclonal primary antibody at 1:50; use PBS or serum as negative control. Detect using the ABC immunohistochemistry method, counterstain with hematoxylin or eosin, and count total and BrdU-positive cells in 10 random high-power fields to calculate the labeling index.
The source describes a scratch-wound model to assess migration of tumor cells in vitro.
Experimental method:
Seed XXX cells at 5–10×10⁵/mL in a 24-well plate, 500 μL per well, in the medium named “RMPI.1640” with 10% fetal bovine serum. Culture 16–24 h to form a monolayer, scratch with a 10 μL pipette tip or sterile toothpick, and wash three times with PBS. The source then says to incubate 24 h, replace with the same 10% serum medium and incubate another 24 h. Remove medium, wash three times with PBS, observe and photograph by inverted fluorescence microscopy. Confirm the medium name and the stated two incubation periods.
The source distinguishes cell–cell and cell–matrix adhesion and describes altered adhesion as relevant to tumor-cell dissemination.
Experimental method:
Coat a 96-well plate with fibronectin at 10 μg/mL, 70 μL per well, overnight at 4°C. Wash three times with PBS, block with 1% BSA at 37°C for 1 h and wash three times. Digest logarithmically growing XXX cells, suspend in serum-free medium at 5×10⁵/mL, and seed 5,000 cells per well in three replicate wells per group. Incubate at 37°C for 1 h and wash away unattached cells with PBS.
Detection method 1:
Fix with 3.5% glutaraldehyde at 4°C for 0.5 h and wash three times with PBS. Stain with 0.1% crystal violet for 0.5 h at room temperature, wash three times, add 100 μL of 10% acetic acid per well and read absorbance at 595 nm after 5–10 min as an indicator of adherent cells.
Detection method 2: Fix with 100 μL methanol per well for 15 min; add 100 μL Giemsa stain per well for 15 min, wash with PBS, and count and photograph adherent cells in five random fields using an inverted microscope.
Detection method 3: Estimate cell number by MTT or CCK-8.
For a Transwell migration assay, the insert is the upper chamber and the surrounding well is the lower chamber, separated by a permeable polycarbonate membrane. The source describes seeding XXX cells in the upper chamber and using membranes of different pore sizes or treatments for co-culture, chemotaxis, migration or invasion studies.
Experimental method:
Warm culture reagents and Transwell chambers at 37°C. Digest logarithmically growing XXX cells, wash once each with PBS and serum-free medium, resuspend and adjust to 2×10⁵/mL. Add 600–800 μL medium with 10% serum to the lower chamber and 100–150 μL cell suspension to the upper chamber; incubate 24 h. Remove the insert, aspirate the upper chamber and fix in a well containing about 800 μL methanol for 30 min at room temperature. Transfer to about 800 μL Giemsa stain for 15–30 min, rinse gently with water several times, and carefully wipe cells from the upper membrane surface with a wet swab. Remove the membrane, air-dry with the lower surface upward, mount with neutral resin and count nine random microscopic fields.
The source describes Transwell invasion as an application of the chamber technique to tumor-cell invasion.
Experimental method:
Matrigel preparation: place Matrigel stored at −80°C at 4°C overnight (the source also says 24 h) to liquefy. Mix 60 μL Matrigel (or 50 μg per chamber as stated) with 300 μL serum-free medium at 4°C; add 100 μL to each of three upper chambers and incubate at 37°C for 4–5 h until a white layer forms. Digest XXX cells, wash three times with serum-free medium and prepare a suspension. Wash the Matrigel once, add 100 μL cell suspension to each upper chamber and 500 μL conditioned medium containing 20% FBS to the lower chamber. Incubate at 37°C for 20–24 h, wash twice with PBS and fix with 5% glutaraldehyde at 4°C. Stain with 0.1% crystal violet or Giemsa, wash twice, wipe cells from the upper surface and count nine random microscopic fields. The source gives both 5–10 min and 0.5 h for staining without distinguishing conditions; verify the intended duration.
The source describes tumor angiogenesis as involving matrix degradation, endothelial migration and proliferation, tube/branch formation and new basement membrane formation. It discusses abnormal tumor vessels as one possible route for tumor-cell entry into circulation and distant dissemination.
Experimental method:
The source instructs placing Matrigel at its stated stock concentration of 10 mg/mL at 4°C overnight, adding 40 μL per well to a 24-well plate and incubating at 37°C for 2 h. Digest and count XXX cells; add 1 mL cell suspension of the chosen density per well in three replicate wells per group. Continue culture at 37°C and photograph at 12, 24, 48, 72 and 96 h under phase contrast at 100–400×. Count tube formations per well using Image-Pro Plus. Confirm the source description of the chilled Matrigel state and the assay timing.
The source describes propidium iodide (PI) as a DNA-binding fluorescent dye excluded by intact cell membranes but entering membrane-damaged cells. It notes use with Calcein-AM or FDA for live/dead staining and gives excitation and emission wavelengths of 535 nm and 615 nm for the PI–DNA complex.
Experimental method:
Wash growing XXX cells with 3 mL PBS and digest with 1 mL trypsin for 1–5 min. Add 5 mL PBS, transfer to a 15 mL tube, centrifuge at 1,500 rpm for 5 min and discard supernatant. Resuspend in 500 μL PBS, add 2 mL of the source’s “20°C 95% cold ethanol” dropwise while vortexing, mix and fix for 30 min. Wash twice with 5 mL PBS and centrifuge at 1,500 rpm for 5 min each time. Resuspend in 800 μL PI stain, incubate in the dark for 30 min at room temperature and analyze by flow cytometry. The ethanol temperature and description “cold” are inconsistent; verify the intended condition.
The source describes phosphatidylserine exposure on the cell surface during early apoptosis and binding by calcium-dependent annexin V (35–36 kDa). FITC-, PE- or biotin-labeled annexin V can be used with flow cytometry or fluorescence microscopy. PI enters membrane-compromised cells; combined staining helps distinguish viable, early-apoptotic and membrane-compromised populations, subject to controls.
Experimental method:
Collect XXX cells in a 10 mL centrifuge tube at the source-stated 1–5×10⁶/mL. Centrifuge at 1,000 r/min for 5 min, discard medium, wash once with incubation buffer and centrifuge again. Resuspend in 100 μL labeling solution and incubate 10–15 min at room temperature in the dark. Centrifuge at 1,000 r/min for 5 min and wash once. Add fluorescent SA-FLOUS solution and incubate at 4°C for 20 min in the dark with occasional agitation. The source specifies 488 nm excitation, a 515 nm band-pass filter for FITC and a filter above 560 nm for PI.
The source interprets FITC−/PI− as viable cells, FITC+/PI− as early-apoptotic cells and FITC+/PI+ as nonviable cells. Its statement that apoptotic cells generally exclude PI should be read in the context of early apoptosis; late apoptotic and necrotic cells may both be PI-positive and require appropriate controls.
The TUNEL method labels 3′-OH ends at DNA breaks in situ using a detectable marker such as fluorescein, allowing observation by fluorescence microscopy.
Experimental method:
Fix → permeabilize → add TUNEL reaction mixture → add converter-POD → add substrate → observe.
Air-dry treated cell coverslips or smears, fix at room temperature for 15–30 min, wash three times in PBS for 5 min each and incubate with blocking solution for 10 min. Rinse and permeabilize for 30 min. The source specifies a treatment mixture of 50 μL TdT plus 450 μL fluorescein-labeled dUTP; the negative control receives only 50 μL labeled dUTP, while the positive control is first treated with 100 μL DNase I at room temperature to 37°C for 10–30 min. Add 50 μL reaction mixture to each dried specimen, cover and incubate in a dark humid chamber at 37°C for 60 min; wash three times with PBS. The source allows fluorescence counting with 450–500 nm excitation and 515–565 nm detection. For POD detection, add 50 μL converter-POD and incubate at 37°C for 30 min, wash three times, add 50–100 μL DAB substrate at 15–25°C for 10 min and wash three times. Photograph, counterstain briefly with hematoxylin or methyl green, rinse immediately, dehydrate through graded alcohol, clear in xylene and mount with neutral resin. The final instruction to add PBS or glycerol after mounting is retained as an ambiguity in the source; verify the intended observation sequence and reagent volumes.
The source describes JC-1 as a cationic dye used to monitor mitochondrial membrane potential. It associates green monomers with lower potential and red aggregates with higher potential, and lists 527 nm and 590 nm as “excitation” wavelengths; verify whether these refer to excitation or emission in the actual reagent instructions.
Experimental method:
Induce apoptosis in XXX cells as appropriate, wash three times with PBS and collect no more than 1×10⁶ cells. Dilute 100 μL 10× incubation buffer with 900 μL sterile deionized water to 1× and prewarm to 37°C. Mix 500 μL 1× buffer with 1 μL JC-1, resuspend cells in 500 μL working solution and incubate at 37°C, 5% CO2 for 15–20 min. Centrifuge at room temperature at 2,000 rpm for 5 min and wash twice with 1× buffer. The source then says to resuspend in 500 μL 10× buffer before flow analysis; verify whether 1× was intended.
The source describes fluorescence-activated cell sorting as measuring scatter and fluorescence of labeled individual cells or particles in flow, then separating selected populations or single cells by instrument settings.
Experimental method:
Perform a preliminary experiment to optimize instrument settings and gates. Based on the proportion of XXX cells, adjust sample concentration and flow so XXX cells pass at 100–300 events/s, preferably around 200. In the historical CellQuest workflow, connect from the Acquire menu, open the saved acquisition window and settings, and load the sample tube. In Sort Setup, choose a gate based on the percentage of XXX cells and the experiment; the source says NoGate acquires without sorting. Select SortCount 0 for continuous sorting and a SingleCell, Recovery or Exclusion mode, and hide aborted-cell counts. Start fluidics and acquisition; when finished, pause and abort, replace the sample with 1 mL distilled water, select standby and remove the collection tube. Centrifuge at 300×g for 5 min, resuspend XXX cells in medium and culture as needed. The source uses the menu spelling “Aquire” in places; verify the instrument software instructions.
Immunomagnetic sorting uses antibodies coupled to magnetic beads to retain cells expressing the target surface antigen in a magnetic field; cells not bound to the beads pass through.
Experimental method:
Collect XXX cells by centrifugation and resuspend at 0.5 mL per 1×10⁸ cells in PBE (0.5% BSA and 0.08% EDTA in pH 7.2 PBS, vacuum-filtered and degassed as stated). Add primary antibody to 10–20 μg/mL and incubate at 4°C for 30 min. Wash with 20 volumes PBE, resuspend at 0.3 mL per 1×10⁸ cells, add secondary-antibody-coated magnetic beads and incubate at 8–15°C for 10–15 min. Place the column in the magnetic field, prewet with 0.5 mL PBE and allow it to drain by gravity. Load the cell suspension, allow it to drain and wash twice with 0.5 mL PBE. Remove the column from the field and flush bound cells with 1–2 mL PBE using the plunger. Wash once in medium.
Cell transfection introduces exogenous molecules such as DNA or RNA into eukaryotic cells for applications including gene-function studies, expression regulation, mutation analysis and protein production.
Experimental method:
Preparing the transfection reagent: add 400 μL nuclease-free water and vortex for 10 seconds to dissolve the lipid material. The source says to store the reagent at −20°C and vortex before use. Select a lipid volume-to-DNA mass ratio of 1:1–1:2, add an appropriate volume of serum-free medium and a suitable amount of MyoD or EGFP DNA to a tube, vortex, add transfection reagent and vortex again. Leave at room temperature for 10–15 min. Remove culture medium, wash once with PBS or serum-free medium, add the mixture and return XXX cells to the incubator for 1 h. Depending on cell type, remove the mixture if appropriate, then add complete medium and culture for 24–48 h. Confirm the reagent-specific storage and mixing instructions.
The historical source describes lentiviral vectors as capable of transducing several cell types, including neurons, hepatocytes, cardiomyocytes, tumor cells and stem cells. Its broad comparative statements about infection and immune responses are not universally established and depend on vector, cell type and conditions.
Experimental method:
Before a larger experiment, optimize cell seeding density, lentiviral amount, puromycin concentration and infection time. Seed XXX cells in a 6 cm dish at an appropriate density with 6 mL medium. For adherent cells, seed XXX cells one day before transduction; for suspension cells, seed XXX cells on the day and include polybrene in the medium. For XXX cells, replace or supplement the medium with polybrene-containing medium to the stated final concentration of 8 μg/mL, add virus and incubate overnight. Change to 6 mL fresh medium after 24 h, including antibiotic if selection is required. The source gives 2–5 μg/mL as a commonly used puromycin range and states that it must be optimized for each cell line. Replace medium every few days as needed; incubation duration depends on the subsequent experiment.
The source distinguishes transient expression, described as nonintegrating and usually lasting days, from stable expression, described as genomic integration and longer-term expression. Stable-cell selection commonly uses the resistance marker on the vector; the source lists hygromycin, neomycin and puromycin resistance. Actual persistence and integration depend on the system.
Experimental method:
G418 (Geneticin) is described as a selection antibiotic for cells carrying a neo resistance marker. To determine a minimum lethal concentration, culture cells in 25 mL flasks with 3 mL medium per flask and test 0, 100, 200, 300, 400, 500, 600 and 700 μg/mL G418, refreshing daily. The source defines the minimum lethal concentration as the lowest dose killing all control cells within two weeks. Passage transfected cells into selection medium at that concentration for about two weeks; after controls die, pick individual resistant colonies into a 24-well plate for expansion.
The source describes two routes to stable lines: selection after lipid transfection, or selection following retroviral or lentiviral transduction.
For the lipid-transfection example, digest and count XXX cells 24 h after transfection and seed a 96-well plate at 2–3 cells per well as stated. After attachment, apply antibiotic selection with time and concentration determined by cell type; the source gives approximately one week for G418 and 2–3 days for puromycin, and an approximate 1% efficiency. Seeding 2–3 cells per well does not by itself establish monoclonality; verify clonality independently.
The source describes using packaging cells, commonly 293 cells, to produce vector, determining titer and transducing target cells. It gives 3–5 days for production depending on the vector and antibiotic selection 1–2 days after target-cell transduction. These are historical example conditions and require system-specific verification.
The source describes co-culture uses including supporting or inducing differentiation, maintaining cell function and viability, regulating proliferation, supporting early embryo development and increasing metabolite yield.
Prepare suspensions of cell A and cell B separately: digest, stop digestion, centrifuge, wash 1–2 times with PBS, resuspend in serum-free medium with BSA and adjust density. Culture the two cell types together at a chosen ratio in the same dish; observe morphology and proliferation microscopically and analyze the results.
In a Transwell system, upper and lower chambers are separated by a permeable polycarbonate membrane. The source describes co-culture, chemotaxis, migration and invasion applications using different membrane pore sizes or treatments. For soluble-factor studies it suggests pores smaller than 3.0 μm, with cell A in the upper chamber and cell B below, to study effects of factors released by cell B on cell A. Whether cells cross depends on cell and membrane properties and should be verified.
The source gives an indirect co-culture example examining whether osteoblast-like cells can induce rabbit bone-marrow stromal stem cells toward an osteogenic fate. Seed rabbit osteoblast-like cells in the lower well and rabbit bone-marrow stromal stem cells on the Transwell membrane as the experimental group. As a control, culture stromal stem cells alone in the insert with basal medium below.
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