How to Count Cells Using a Hemocytometer | Protocol & Calculations

This comprehensive workflow demonstrates solving a cell seeding problem step-by-step. Given a six-well plate with 2 mL per well and target of 250,000 cells per well: (1) Convert target to cells/mL: 250,000 × 2 = 500,000 cells/mL; (2) Calculate total volume for six wells plus 10% pipetting error: 6 × 2 + 0.2 = 13 mL; (3) Solve for V1 using C1V1 = C2V2: V1 = (500,000 × 13) / 1,000,000 = 6.5 mL; (4) Calculate fresh media: 13 - 6.5 = 6.5 mL. This systematic approach ensures accurate cell distribution across all wells at the desired density.

This integrated workflow covers post-counting procedures for cell culture passaging. Key steps include: (1) Using a hemocytometer with 10 microliter samples to count cells under microscope; (2) Calculating required volumes to achieve target cell densities (typically 1 million cells per dish); (3) Inverting tubes 10 times to mix settled cells; (4) Transferring calculated cell volumes to fresh media; (5) Labeling new dishes with initials, cell line, and date; (6) Adding complete cell-media mixture to dishes while avoiding bubbles; (7) Swirling to distribute cells evenly; (8) Placing dishes in tissue culture incubator. This systematic approach ensures reproducible cell culture results.

Cell seeding is the process of distributing a specific number of cells onto culture plates to ensure experimental consistency, where cells must be counted using a hemocytometer with trypan blue staining (which stains dead cells blue while healthy cells remain transparent) and then calculated using the formula: Cell Count = (Cells counted × Dilution factor × 10^4) / Volume factor, to achieve the desired cell density for experiments.

This comprehensive workflow demonstrates how to perform automated cell counting using ImagePro software. The process begins with loading a two-channel multi-well plate dataset and accessing the protocol browser to select the Cell Count protocol from the Cell Biology collection. Key steps include synchronizing the protocol with dataset channels, selecting the appropriate counting channel (DAPI), and configuring segmentation parameters. Users learn to select representative objects and background regions, set minimum area filters to exclude noise, and use reference size for clumped cells. The workflow concludes with configuring result displays including heat maps and data collectors, then executing the protocol on the entire plate to generate cell counts for each well.

The complete cryopreservation protocol involves several critical steps: (1) Harvest cells during log phase when viability exceeds 90%, (2) Resuspend cells in serum-containing media, (3) Centrifuge to remove debris and resuspend in 10% DMSO in FCS, (4) Transfer to cryovials, (5) Freeze at -80°C first, then transfer to liquid nitrogen at -196°C. DMSO acts as a cryoprotectant by preventing intracellular ice crystal formation that would otherwise damage and kill cells. The two-stage freezing process ensures gradual temperature reduction, protecting cellular integrity during preservation.

Cryopreservation involves spinning cells down, aspirating medium, and resuspending in 50/50 cell banking medium and growth medium. Final volume per vial is 500 microliters, requiring 1.25ml of each component for 5 vials. Vials are placed on ice briefly (freezing medium can be damaging) then transferred to -80°C freezer for long-term storage. The complete process took approximately 77 minutes from start to finish.

The cryopreservation process involves storing cells in cryovials at -80°C for days before transferring to liquid nitrogen at -196°C. Three protocols exist: 5% DMSO with 95% complete media, 10% DMSO with 90% complete media, and 5% DMSO with 95% FBS (highest recovery rate). Media must be trypsin-free. Rapid thawing at 37°C prevents intracellular ice formation. Applications include cell/organ cryosurgery, blood transfusion, bone marrow transplantation, and IVF.

Proper laboratory cryopreservation requires specific conditions. Cells should be at confluence, healthy, and have viability greater than 80%, ideally at the log phase of their growth curve. The procedure involves adding cryoprotectant media containing DMSO and serum, then transferring cells to containers designed for gradual temperature reduction. Cells are first cooled to -80°C before being transferred to liquid nitrogen storage at approximately -196°C.

This extensive section covers complete cell cryopreservation methodology. Cell freezing creates permanent cell banks for future experiments by preserving genetically identical populations. The freezing medium contains 10% dimethyl sulfoxide (DMSO) in culture medium, prepared by combining DMSO solution with culture medium. DMSO must be added drop by drop over five minutes while keeping cells on ice to prevent osmotic shock. This gradual addition creates a two-fold dilution effect, resulting in the optimal final DMSO concentration of 10%. The complete protocol includes centrifuging cells to form a pellet, resuspending in half the necessary medium, adding freezing medium drop by drop, distributing to cryovials, storing overnight at -80°C, and transferring to liquid nitrogen the next day.

Manual cell counting using a hemocytometer is presented as inefficient and outdated, with automated cell counters offering a superior alternative. The video highlights three key advantages of adopting an automated cell counter: time savings, improved accuracy, and affordability. Users counting five slides per day manually can save approximately 10 hours per month by switching to an automated system with a reusable slide, and up to 15 hours per month when using disposable slides. Automated counters are documented in multiple scientific articles to provide greater accuracy than manual methods, leading to higher-quality lab results. The Countess II FL Automated Cell Counter is specifically promoted as a cost-effective option, being reasonably priced compared to other automated systems and eliminating ongoing slide costs due to its reusable slide design. This positions the device as a one-time investment rather than a recurring expense. The video encourages viewers to abandon manual counting practices and join a broader movement against hemocytometer use. Links are provided to compare automated cell counters, learn about the accuracy and precision of the Countess II FL, and view a tutorial on performing brightfield cell counts. The promotion emphasizes that cost is no longer a valid barrier to automation, urging laboratories to upgrade for efficiency and reliability.

Two primary methods exist for cell counting: manual hemocytometer and automated systems. Hemocytometers are glass slides with etched grids creating defined volumes; cells migrate via capillary action, and counts are multiplied by dilution and volume factors. Automated counters like Countess use plastic slides with imaging technology to automatically detect and count cells, distinguishing live from dead cells and providing corrected viable concentrations. Both methods require proper dilution correction (2x for trypan blue), though automated systems handle calculations internally.

The hemocytometer is the equipment used for complete blood counts. Two main methodologies exist: flow cytometry and electrical impedance. The impedance method, developed by Wallace Couter in 1956, is the primary method in modern analyzers. Cells are counted based on electrical impulses generated when immersed in a conductive medium. This disturbance in the electric field helps identify different cell types including leukocytes, erythrocytes, monocytes, and lymphocytes.

Automated cell counters (Sysmex, BD, Coulter) are highly accurate when properly maintained with daily quality control. They count cells using laser technology and electrical impedance. Manual techniques are needed for: (1) WBC differential and morphology; (2) Giant platelets; (3) Platelet clumps; (4) When automated results are unreliable. Proper peripheral smear preparation requires good staining technique.

Automated cell counters perform complete blood counts using three principles: electrical impedance (detects cell size via current disruption), light scatter/flow cytometry (forward scatter indicates size, side scatter indicates complexity), and image-based analysis (microscopy with statistical algorithms). Operation modes include manual aspiration and automated tube loading. Differential counting uses three-part analyzers (lymphocytes, mixed population, granulocytes) or five-part analyzers (neutrophils, lymphocytes, monocytes, eosinophils, basophils). Five-part analyzers combine impedance, light scatter, and flow cytometry to identify all WBC types separately. Some analyzers count nucleated RBCs and flag abnormal cells.