Lab Descriptions

Instructors: Mark Naivar, Jim Freyer & Ben Finnin 

The purpose of this laboratory is to discuss instrumental and operational parameters that can affect the quality of the data collected from a flow cytometer. While specific examples will be used to illustrate the concepts, the goal is to provide the participants with tools and information that can be applied to a wide variety of biological applications. 

Instructors: Kathleen McGrath & Aja Rieger

 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 how to analyze data that contain both types of information to answer biologically relevant questions. In this lab, we will be predominately be utilizing and analyzing data from ImageStream flow cytometer (ISXMarkII) from Cytek (Amnis Corp). However, we will also compare the imaging analysis in expanding options integrating imaging in flow cytometry including BD FACSDiscover S8 Cell Sorter and ThermoFisher Attune CytPix Flow Cytometer.

The bulk of the time, the class will focus on independent learning hands-on tutorials based on real-life examples that will teach fundamentals and advanced skills in the combined gating of intensity level, shape, size, and texture to address a specific biological questions. 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 beginner level to challenging experienced imaging flow cytometrists, with the central concepts of gating, masking, and combined morphometric/fluorescent feature selection using machine learning to masks combined features. Students will also run the ImageStream and look “under the hood” to see how it works. We will also cover best practices for publishing imaging flow data and how to encourage and train new users. 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.

Instructors: Fred Preffer & 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 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 discuss the mechanics of staining (surface versus intracellular) as well as all the reagents necessary to properly develop successful staining. Participants will perform viability staining, blocking, antibody staining, fix/perm, intranuclear staining and sample acquisition while learning best practices. Provisionally, we are planning on the analysis of 10 colors plus a live/dead marker e.g.   CD3, CD4, CD8, CD14, CD16, CD19, CD25, CD45, CD56 and FoxP3.  What is learned from this panel will be transferrable to any desired panel the participants wish to stain in their own labs. 

Instructors: Patricia Rogers, Jennifer Couget & Si-Han Hai 

Instructor: Joe Tario, Jenna Hope & Kathy Muirhead 

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?

Instructor: Paul Wallace, Kah Teong Soh & Vince Shankey 

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.

Instructors: Vera Tang, Josh Welsh & Joanne Lannigan 

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.

Instructors: Alex Henkel & Zach Stenerson 

Instructors: Riu Gardner & Alan Saluk 

The evolution of high-speed droplet-based cell sorting instrumentation has developed over the decades to allow ease of access to contemporary biological researchers.  While the instrumentation has become much more user friendly than before, there is an art to mastering this unique intersection of biology and physics.  We will work with students to break-down these overlapping factors with hands-on demonstrations that will attempt to demystify many of the factors that allow for successful sorting of various experimental types. 

From basic stream physics and optimized sorting parameters to sample preparation optimizations, we will address the students based on their skill levels with hands-on sorting with a modern high-speed cell sorter.  The goal of the course is for users to be able to build on their existing knowledge base to develop a deeper more nuanced understanding of what makes for successful for experimental designs for cell sorting.

Instructors: Jonathan Irish & 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 may not be sufficient. Computational techniques have been developed to analyze, visualize, and interpret these data. These techniques fall into seven primary categories: automated quality assessments, dimensionality reduction, automated cell and sample classification, normalization and batch effect removal, differential statistics, and visualization.

This lab will introduce attendees to some of these techniques and approaches. We will discuss when, why and how to use algorithmic approaches with some discussion on theory and a lot of hands-on application using high-dimensional data. We will use FlowJo™ version 10.10, web based tools, and R to walk through performing high-dimensional workflows, focusing our discussion on when an algorithm can expedite your analysis, how to use it, and what the differences are in similar algorithms, for example, tSNE and UMAP.  Our goal is to give you a good understanding of the mechanisms of a wide range of algorithms and the ability to use them in available software.

Instructors: Lola Martinez & Anna Belkina

This laboratory will use different spectral flow cytometers from different manufacturers (BD, Cytek and Sony) to learn key concepts to take into consideration when troubleshooting your spectral flow experiment.

Together, we will cover settings optimization, threshold, area scaling factor, importance of reagent titration and good controls to succeed.

We will go over different data sets covering multicolor immunophenotyping and high autofluorescence samples expressing fluorescent proteins to discuss how and when to carry autofluorescence removal in our samples.

A short survey will be sent before the course to assess students’ experience in spectral cytometry. We will work with a small group of students, and we will try to group them by their previous experience, trying as much as possible to tailor the sessions to the experience and cytometer access of each group.

Instructors: Rachel Sheridan & Kathryn Fox

In this lab, you will stain PBMCs with antibodies for a small panel of surface markers and viability dyes. You’ll titrate the antibodies and explore variables such as cell number, antibody concentration, and reaction volume. Together, we will interpret the results of various staining conditions, and discuss key factors in staining cells with antibodies and viability dyes. 

Instructors: Yoav Altman & Cheryl Kim

Flow cytometers require regular maintenance and QC to ensure consistent performance and data reproducibility over time.  Diagnosing and troubleshooting instrument problems are critical skills for personnel in charge of flow cytometers. In this hands-on lab, we will review key operating principles, instrument standardization, and maintenance routines.  We will then take a deep dive into how problems in the fluidics, optical and electronic systems can be diagnosed using a variety of tools including commercially available QC beads, manufacturer recommended procedures and instrument QC reports. Upon completion of this lab session, students will have newfound confidence overseeing and operating instruments, and performing minor repairs of malfunctioning cytometers.