Lab Descriptions

General Lab Descriptions

*Labs are subject to change*

1. Build a Flow Cytometer

Instructors: John Martin, Mark Wilder, Travis Woods

This lab is designed to give the students a better appreciation of the inner workings of a flow cytometer, taking away some of the mysteries of what is hidden inside the cabinet.

In this lab session you will assemble a small flow cytometer and use it to measure a sample of fluorescent microspheres. You will learn some of the important steps used in setting up a flow cytometer, some diagnostic clues that are useful for evaluating a flow cytometer’s performance and an appreciation for what is involved in constructing flow cytometers.

This compact flow cytometer is assembled using modular parts according to a detailed protocol. It includes the following components: laser, laser beam shutter, laser power attenuator, beam block, CCD camera and video monitor (used for viewing the laser beam – sample stream intersection region), flow chamber, fluorescence collection optics, fluorescence detector, electronics, oscilloscope and a computer.

Because you will be installing and aligning optical components along a laser beam line good laser safety practice will be discussed and stressed. By the end of the lab session, you will have assembled a working cytometer and will be analyzing the microsphere sample and optimizing the final adjustments to obtain the best CV. In recent years, CVs below 2% have been achieved.

2. Flow Cytometry Data Acquisition

Instructors: Mark Naivar and Jim Freyer

This laboratory will use flow cytometer simulators to learn key concepts of data acquisition, and also cover some basic data analysis. Regardless of whether you are going to gate or model your data to get your final result, and regardless of which instrument you are using, it is important to acquire flow data as accurately as possible. It is much better to collect your data properly up front, rather than try to “fix” poor data after the fact (many acquisition problems simply cannot be “fixed”).

Small teams of students will use flow cytometer simulators to explore critical factors that affect data acquisition (sample preparation, thresholding, PMT voltage, coincidence, aggregates, and sample throughput). Students will be able to reconfigure the cytometer simulator in ways that are impossible for real instruments in order to provide a more intuitive feel for how different conditions can affect data acquisition. We will also cover the basics of data visualization and gating, which are important for evaluating and adjusting acquisition parameters. Finally, the simulator will be used to explore some basic examples of compensation to give students a much better feel for when compensation is important, why it is needed, and how it improves the data that are collected. Because we will be using a simulator, no cells will be harmed, and no one will run out of sample! No lab coats, gloves or goggles required!

3. A Typical Cytometry Day

Instructors: Kathryn Fox and Rachael Sheridan

You have a big flow experiment to run today. What can you do to collect high quality data? We’ll walk through flow cytometry best practices in sample preparation, controls, instrument set up and QC, and data acquisition. We’ll demystify the cytometer ‘black box’ through discussion of how flow cytometers work and use hands-on activities to show how these principles act in practice to contribute to data quality. Bring your questions and experience a day in the flow lab!

4. Next Generation Cell Sorting: New Tools and Methods

Instructors: Rui Gardner and Zach Stenerson

This practical laboratory session will focus on several areas of interest in cell sorting that apply to particle sorting in general. We will cover instrument setup based on the task at hand. In other words, how to realistically approach optimizing nozzle size, stream stability, deflection envelope, break off, drop rate and sample rate for any given experiment. The lab will try to provide the attendee with approaches for use in their own facility in problem solving a wide variety of sorting experiments, regardless of the cytometer they use, including suggestions on advising facility users on sample preparation.

5. Integrating Flow and Imaging Cytometric Analyses

Instructors: Kathleen McGrath and Aja Reiger

Flow cytometry and microscopy are two powerhouse single cell analytical technologies. When combined, they can give unprecedented information about cell biology. In this lab, you will learn to how to analyze data that contain both types of information to answer biologically relevant questions. We will be utilizing an ImageStream flow cytometer (ISXMarkII) from Luminex (Amnis Corp). The bulk of the time, the class will doing independent hands-on tutorials of your choice that will teach fundamentals and advanced nuances in the combined gating of intensity level, shape, size, and texture to address a specific question in that data set. We will be available to assist and answer any questions, but the best way to learn this type of analysis is to do it! These tutorials range from those for training beginners to challenging experienced imaging flow cytometrists, with the central concepts of gating, masking, and combined morphometric/fluorescent feature selection that are core to any flow cytometric imaging analysis. We will include training data sets for the new machine learning module which makes combined features (like PCA or tSNE). Students will also run the ImageStream and look “under the hood” to see how it works. Finally, we will take some short breaks to discuss best practices for publishing imaging flow data, issues and value of compensation in imaging data, and how to encourage and train users in a core facility with this approach. The ultimate goal of the lab is to empower the participants to know if you can see a difference in cells, you can use imaging flow cytometry to quantitate it.

6. Multicolor Immunophenotyping

Instructor: Lauren Nettenstrom

Flow cytometry is a method for analyzing cells for multiple surface and intracellular proteins utilizing excitation lasers and monoclonal antibodies conjugated to unique fluorescent tags. Additionally, simultaneous light scatter measurements that impart cell size and complexity are coupled with this information to identify and describe individual leukocyte cell populations. There will be a concentrated discussion about the antibody/antigen reaction, antibody kinetics, antibody titration, conjugated versus unconjugated antibodies, the importance of choosing the correct monoclonal antibody clone as well as choosing the correct antibody/fluorochrome combination. We will go through the steps of successful panel design, perform an antibody titration, calculate the antibody’s signal-to-noise ratio, stain different combinations of reagents, determine both the optimal antibody clone and its preferred fluorochrome combination, and discuss the mechanics of staining, surface versus intracellular as well as all the reagents necessary to properly develop successful staining.

7. Optimizing and Troubleshooting Spectral Unmixing

Instructor: Laura Johnston

Spectral flow cytometers use the same operational principles as conventional flow cytometers with the exception of the way that fluorescence is collected. Instead of capturing the peak emission using a dedicated optical filter, the full emission of each fluorochrome is measured across a detector array to generate a fluorescence signature. Spectral unmixing is a mathematical algorithm that uses the signatures provided by single stained controls to distinguish multiple fluorophores within a multicolor sample. In this lab, participants will work with spectral data sets to learn a basic workflow for unmixing. This includes choosing appropriate controls, setting up the unmixing wizard, and troubleshooting basic unmixing errors. The autofluorescence extraction feature and its impact on unmixing will also be discussed. By the end of this lab, participants should be able to identify and avoid common mistakes in unmixing. 

8. Single Cell Genomics

Instructors: Patricia Rogers and Jennifer Couget

This lab will give students an understanding of the different commercially available single cell technologies on the market and go into details of what applications they are used for (10x Genomics, Mission Bio, BD Rhapsody — to cite a few but not limited to). We will go through the workflows by starting with practical aspects of cell enrichment and viability analysis to hands on practice with loading samples in order to create single cell emulsions. From there we will go through library construction basics and which instruments will be used for quality control assessments. Finally, students will get an understanding of failure points during workflows and how to troubleshoot when libraries fail to generate good sequence results.

9. Tracking Immune Responses: Antigen Binding, Proliferation, & Apoptosis

Instructors: Kathy Muirhead and Joe Tario

Flow cytometry is a powerful tool for monitoring three key aspect of adaptive immunity: antigen recognition, clonal expansion, and apoptotic cell death. In this lab we will focus on critical issues for:

  • Identification of antigen-specific T cells using multimer labeling.
  • Proliferation monitoring using dye dilution.
  • Early apoptosis detection using probes for caspase activation, membrane integrity, and phosphatidylserine externalization. 

Participants will be divided into small groups for hands-on experience with:

  • Immunophenotypic characterization of low-frequency multimer binding T cells.
  • Staining optimization for proliferation analysis using protein-reactive or membrane intercalating dyes (CFSE, PKH26 and newer analogs of each).
  • Probe selection for multicolor assays correlating antigen specificity with downstream outcomes (e.g., cell division, apoptosis).

We will also cover several instrument setup and data collection issues likely to be of interest even if your laboratory is not already doing functional assays, including:

  • Color compensation – recognizing problems, optimizing probe combinations to minimize them.
  • Collection and analysis of data on low frequency subpopulations – are these events real or are they junk?
10. Intracellular Cytometry: Cell Signaling, mRNA Transcription, & Cytokine Synthesis

Instructors: Paul Wallace, Vince Shankey, Kah Teong Soh

Lipopolysaccharide (LPS), a cell wall component found in most Gram-negative bacteria, activates signaling cascades in several different cell types, including some hematopoietic cells. Monocytes express surface Toll-like receptor-4 (TLR4), which binds LPS, and in conjunction with CD14, initiates a signaling cascade that results in the degradation of IkB (Inhibitor of NFKB) and release of NFKB (nuclear factor kappa B), with the latter localizing into the nucleus. There, NFKB binds to transcriptional initiation sites, resulting in the synthesis of new mRNA and translation into monokines and other proteins. Given the complexity of the response triggered by different cell populations, single cell flow cytometric analysis of signaling and downstream protein/cytokine expression patterns has greatly advanced our understanding of the immune system. 

This lab will follow the kinetics of antigen binding, NFKB translocation, mRNA transcription, and protein expression. We will:

  • Review the signal transduction pathways that regulate the acute inflammatory response via the NFKB transcription factor and methods to measure them.
  • Discuss technical variations to simultaneously detect cell surface and intracellular targets.
  • Present a simple approach to fixation and permeabilization which provide access to cytoplasmic and nuclear compartments.
  • Discuss the technical aspects of simultaneously measuring intracellular mRNA and protein targets.
  • Understand and appreciate the cytokine/monokine mRNA and protein kinetic profiles in activated lymphocytes and monocytes.
  • In the hands-on wet lab portion, participates will stain and analyze control and activated lymphocytes for surface markers (CD45, CD3, CD4, and CD8), intracellular cytokines (IFNγ, IL-4, and IL-2), and viability.
11. Flow Cytometry Analysis of Extracellular Vesicles

Instructors: Vera Tang and Joshua Welsh

While implementing best practices is important in all cytometry, it is especially important when analyzing extracellular vesicles and other small particles because of their size. In this lab, you will learn how to optimize and quantify the sensitivity of your instrument for detecting small particles, determine whether you are detecting single particles or “coincidence”, utilize controls to interpret results, and calibrate your fluorescence and light scatter signals for downstream analysis and reporting in standard units.

12. Panel Design and Optimization

Instructors: Dagna Sheerar and Alex Henkel

In this lab students will walk through the workflow of designing an optimized, rigorous, and reproducible flow cytometry panel. The tips and tricks shared in this lab will be applicable to all panels, small or large. With the goal of maximizing sensitivity for each marker detected in the assay, we’ll discuss power calculations to determine sample size, sample preparation, protocol development and optimization, annotation and record keeping, characteristics of available fluorochromes, characteristics of markers of interest, how to optimally pair markers with fluorochromes, proper controls, instrument characteristics, spillover spreading, assay standardization, and considerations for high dimensional data analysis. Students attending this lab should walk away with an understanding of proper considerations and a workflow for designing an optimal, rigorous, and reproducible flow cytometry assay.

13. High Dimensional Data Analysis

Instructors: Beth Hill and John Quinn

Recent advancements in cytometry allow us to measure an increasing number of features per cell, generating huge high-dimensional datasets. This creates challenges for data analysis. Traditional approaches based on subjective manual gating on biaxial plots are not sufficient.  Computational techniques have been developed to analyze, visualize, and interpret these data.   These techniques fall into five primary categories: automated quality assessments, dimensionality reduction, automated cell and sample classification, normalization and batch effect removal, and visualization.

This lab will introduce attendees to some of these techniques and approaches. We will discuss the pros and cons and practical aspects of different methods using a hands-on manner with a high-dimensional dataset. You will learn what PCA, t-SNE, UMAP and Cen-se’ can do for you and how they differ. We will develop an automated analysis strategy using probability state modeling and GemStone™ 2.0. In addition, attendees will create a high-dimensional workflow in FlowJo™ 10.8. Finally, we will discuss other algorithmic approaches and options for high-dimensional analysis.

14. Cytometric Analysis and Sorting of Plants, Algae, Cellular Homogenates and Large Particles

Instructors: David Galbraith and Claire Sanders

This laboratory provides practical strategies for handling “unusual” samples in flow cytometry and cell sorting, including dealing with large cells and other biological particles, and with non-mammalian species. Flow cytometry is ideally suited for measurement of single cell suspensions, and the standard designs of flow cytometers and cell sorters originally accommodated the typical size range of blood cells, with diameters spanning ~10-20 µm. However, across the domains of life, cells are found that fall outside of this range. Many eukaryotic cells are larger than 20 µm, and sometimes much larger. Correspondingly, examples of algae and photosynthetic prokarya are increasingly identified that are smaller than 10 µm. Further complications are introduced when we encounter organisms, or their tissues, that are not cellular suspensions. Converting tissues and organs to single cells is not necessarily a simple task, since it requires dissolution of the extracellular matrices that interlink the cells without otherwise perturbing the cells. In this lab, we will demonstrate (a) analysis and sorting of cells that are either larger or much smaller than the mammalian cells typically encountered in the shared user flow core, (b) how simple homogenization techniques can be used to release suspensions of subcellular organelles, particularly nuclei, for flow analysis and sorting, (c) how to analyze microalgae using flow cytometry, and how to monitor changes in cellular parameters during culture.

All flow operators should benefit from the materials covered in this lab, since inevitably they will be encounter users that wish to analyze and sort unusual samples.

As part of this lab, we will also be demonstrating operation of the Velocyt cytometer from BennuBio. This sheathless high throughput analyzer is capable of analyzing particles up to 1 mm and recovers the full undiluted sample.