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Measuring relative lysosomal volume for monitoring lysosomal storage diseases
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Danielle te Vruchte1, Kerri L. Wallom, Frances M. Platt1 1
Department of Pharmacology, University of Oxford, Oxford, UK
Corresponding authors: E-mail: [email protected]; [email protected]
CHAPTER OUTLINE 1. 2. 3. 4.
Measuring Relative Lysosomal Volume as an Index of Lysosomal Storage .............. 332 In Which Circulating Cell Type Should Relative Lysosomal Volume Be Measured?.. 333 How to Measure Relative Lysosomal Volume in Blood Cells .................................. 334 Methods ............................................................................................................ 336 4.1 Preparation of Human Blood ................................................................ 336 4.1.1 Materials and reagents ..................................................................... 336 4.1.2 Protocol ........................................................................................... 337 4.2 Preparation of Cells from Mouse Spleen or Whole Blood ......................... 337 4.2.1 Materials and reagents ..................................................................... 337 4.2.2 Protocol ........................................................................................... 338 4.3 B-Lymphocyte Staining........................................................................ 339 4.3.1 Materials and reagents for human cells ............................................. 339 4.3.2 Protocol for human cells ................................................................... 339 4.3.3 Materials and reagents for mouse cells ............................................. 339 4.3.4 Protocol for mouse cells.................................................................... 340 4.4 Flow Cytometry, Calibration, Acquisition, and Analysis ........................... 340 4.4.1 Materials and reagents ..................................................................... 340 4.4.2 Protocol ........................................................................................... 340 4.5 Isolation of B-Cells for Biochemical Assays or Microscopy....................... 342 4.5.1 Materials and reagents for human cells ............................................. 342 4.5.2 Protocol for human cells ................................................................... 342 4.5.3 Materials and reagents for mouse cells ............................................. 342 4.5.4 Protocol for mouse cells.................................................................... 343 4.6 Influence of Patient Blood Shipping Times ............................................ 343 4.7 Influence of Blood Storage Temperature on LysoTracker Staining ............ 344
Methods in Cell Biology, Volume 126, ISSN 0091-679X, http://dx.doi.org/10.1016/bs.mcb.2014.10.027 © 2015 Elsevier Inc. All rights reserved.
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4.8 Influence of Delays in Analysis of Samples Post-LysoTracker Staining...... 345 4.8.1 Some economic considerations relating to B-cell staining .................. 345 5. Summary ........................................................................................................... 346 Acknowledgments ................................................................................................... 346 References ............................................................................................................. 347
Abstract Biomarkers are important tools in medicine, which can be used for monitoring disease progression and response to therapy. One of the main problems in rare lysosomal storage diseases is that there are over 70 different diseases, all with different biochemical storage profiles. Developing biochemical biomarkers therefore requires an individual assay per disease/subgroup of diseases. An alternative approach is to develop an assay that is independent of the specific macromolecules stored. This chapter discusses an assay that may serve as a universal biomarker for these diseases and measures the expansion of the late endosomal/lysosomal system. We have developed an assay that takes advantage of a commercially available late endosomal/lysosomal probe, LysoTracker, which becomes trapped in the acidic compartment of cells and emits a fluorescent signal that can be detected using flow cytometry. In this chapter, we detail the methodology behind this assay and discuss the factors that need to be considered when establishing this assay in clinical and research settings.
1. MEASURING RELATIVE LYSOSOMAL VOLUME AS AN INDEX OF LYSOSOMAL STORAGE The late endocytic system comprises two main acidic organelles, namely late endosomes (LE) and lysosomes (Lys). Their volume is tightly regulated in a cell-typespecific manner. Resting cells tend to have a lower LE/Lys volume compared to highly active cells (e.g., macrophages). The function of LE/Lys is the catabolism and recycling of cellular macromolecules (Luzio, Pryor, & Bright, 2007). However, in addition they serve as nutrient sensors (Zoncu et al., 2011) and are signaling organelles in part through their role as regulated calcium stores (Churchill et al., 2002; Calcraft et al., 2009). Irrespective of the cell type, if there is an inborn error of metabolism that affects lysosomal function, substrates and cargos within LE/Lys build up leading to “storage” and disease (Platt, Boland, & van der Spoel, 2012). Typically, these inherited diseases are caused by mutations in genes encoding lysosomal enzymes (acid hydrolases), but defects in non-enzymatic lysosomal proteins (soluble or membrane proteins) can also lead to lysosomal storage (Winchester, Vellodi, & Young, 2000; Wraith, 2002; Beutler, 2006). The accumulation of non-degraded metabolites in the LE/Lys system then triggers an increase in LE/Lys volume, providing a potentially universal cellular biomarker for these diseases (Vitner, Platt, & Futerman, 2010). The lysosomal storage diseases (LSDs) comprise >70 individually rare diseases, but are collectively much more common and affect approximately 1:5000 live births
2. Measuring relative lysosomal volume as an index of storage
(Vitner et al. 2010; Platt et al. 2012). The technical advantage of being able to measure relative changes in acidic compartment volume is that it allows storage levels to be measured in patient cells without the need to measure disease-specific biochemical metabolites. Therapies for these diseases aim to reduce LE/Lys storage; therefore, measuring relative lysosomal volume can also potentially report on response to treatment. The current approved therapies for LSDs are enzyme replacement therapy, bone marrow transplantation, and substrate reduction therapy using small molecules (Platt et al. 2012; Platt, 2014). However, other therapeutics are undergoing clinical trials at the current time (gene therapy, chaperone therapies, etc.) and have recently been reviewed elsewhere (Platt, 2014). The main practical applications of measuring LE/Lys volume as a potentially universal cellular biomarker for LSDs can be summarized as follows: 1. Suspicion of an LSD: Based on clinical presentation, this can be confirmed at the cellular level in a blood sample or in skin fibroblasts by measuring LE/Lys volume. This can be performed prior to commissioning more costly diseasespecific diagnostic tests, typically enzyme assays and mutation analysis. 2. Following diagnosis of an LSD, peripheral blood cells from the patient can be measured over time, allowing longitudinal progression of storage to be measured quantitatively. 3. In clinical trials, the use of this biomarker may aid stratification of patients for recruitment to trials and also for monitoring individual patient responses to a clinical intervention. 4. In LSDs with approved therapies, relative lysosomal volume can also be used as a monitoring tool to study long-term stability and detect any signs of a failure in treatment efficacy over time. 5. As many LSDs may benefit from combination therapies, in the future this could also be a useful biomarker for determining whether the addition of a new therapeutic agent significantly impacts levels of storage. As blood is a minimally invasive source of patient cells, it is an ideal starting point for such an assay. However, blood contains a great variety of cells of hematopoietic origin, and a very important question is therefore: Which circulating nucleated cell type should be monitored? A second question is: How can relative lysosomal volume be measured in blood cells? We will discuss both of these questions in more detail before discussing the detailed methods.
2. IN WHICH CIRCULATING CELL TYPE SHOULD RELATIVE LYSOSOMAL VOLUME BE MEASURED? What proved to be the most important factor in the reliability and reproducibility of the assay we have developed was the choice of cell type to analyze (Lachmann et al., 2004; te Vruchte et al., 2014). All mononuclear cells have LE/Lys compartments, so are all candidates to be measured? Readers may be thinking why not simply profile
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all mononuclear cells in human blood and take an average or get data on multiple cell types/populations. The reason this is not a good approach is that the hematopoietic system varies significantly with age (a newborn does not have the same circulating blood cell ratios as an infant or an adult) and most significantly these cells change with disease progression in LSDs (Gadola et al., 2006; Speak et al., 2014). Therefore, a specific cell type needs selecting for LE/Lys volume determination to ensure reliable longitudinal analysis in the same individual and permit valid comparisons to be made between individuals. We originally focused on cells with the largest LE/Lys volume (myeloid cells), but they can only be accurately identified using multiple cell surface markers. Very significantly, they change in response to infection to a variety of different activation states, requiring even more markers to be analyzed in order to identify them accurately. It is therefore difficult to be sure that precisely the same cell type is being monitored over time making them a poor choice for this cellular assay. In the end, we focused our attention on small circulating resting B-cells and they indeed proved ideal for our purposes (Lachmann et al. 2004; te Vruchte et al. 2014). Small resting B-cells are antigen naı¨ve and have a very low cytoplasmic-to-nuclear volume. Their acidic compartment volume is therefore relatively low compared with myeloid cells, but they have the major advantage that they all express the pan-B-cell marker CD19. This means that they can be reliably identified with a single antibody without the need to sort or purify the cells, and they are compatible with longitudinal analysis, as they are a homogeneous population that crucially does not change in response to infection.
3. HOW TO MEASURE RELATIVE LYSOSOMAL VOLUME IN BLOOD CELLS There are at least three main ways that relative lysosomal volume could be measured in mononuclear cells: (1) by measuring a lysosomal protein by western blot or enzymelinked immunosorbent assay (ELISA) as a reporter on relative compartment size/ volume, (2) by measuring a fluorescent probe that is endocytosed and “chased” into the late endocytic system, or (3) by using a cell-permeant fluorescent probe that is rapidly trapped in acidic organelles and can be measured to quantify relative volume of LE/Lys. All of these approaches are equally valid but have a number of pros and cons from a practical perspective. Measuring lysosomal proteins by western blotting assumes that the level of the lysosomal protein of interest scales with LE/Lys volume in a given LSD. This method requires the isolation of the cell type to be measured free of contaminating cells; the proteins then need extracting with detergent containing buffer, running on an sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) gel, transferring to a suitable membrane, and probing with an appropriate antibody to detect the lysosomal protein of interest. This is a time-consuming exercise and low throughput. It is also relatively expensive due to the need to run gels, to blot
3. How to measure relative lysosomal volume in blood cells
and probe the blot with a reasonable volume of antibody. ELISAs have a similar limitation in that the cells need isolating to high purity, but in this case there is no need to run gels, as the lysosomal protein is simply measured in an ELISA plate. Of the two methods, ELISAs are higher throughput, but the cell isolation step is the big drawback. Even a small contamination with myeloid cells can distort the lysosomal marker levels in a B-cell preparation very significantly. However, the advantage of these approaches is that they can be performed on frozen material, as long as the cell purification is performed on fresh cells. The alternative is to pulse the cells in question with a fluorescent reporter that is “chased” into LE/Lys. Fluorescent dextran, for example, could be used for this purpose (Bright, Gratian, & Luzio, 2005; Lloyd-Evans et al., 2008). However, pulsechase times need to be optimized and the chase time significantly extends the assay time. Again, making sure a single homogeneous cell population is studied is crucial, as the chase times will be cell-type dependent. This requires live cell analysis by microscopy, so they must be analyzed before viability is compromised. The analysis is tedious and not particularly quantitative. The method we have adopted and optimized uses a cell-permeant dye, such as LysoTracker, which is a weak amine. At acidic pH, it is protonated and becomes trapped in LE/Lys. The relative fluorescence of the signal is then proportional to the relative volume of the LE/Lys compartment (te Vruchte et al., 2014). There are several advantages of this method. Firstly, it can be coupled with very sensitive analytical flow cytometry that has a large dynamic range. B-cells are simply identified in the circulating mononuclear cell populations with an anti-CD19 antibody (e.g., red conjugate). The cells are then co-labeled with LysoTracker (e.g., LysoTracker green) and the B-cell population analyzed without the need to sort or purify B-cells using other methods. This greatly speeds up the assay. The second advantage is that the antibody-staining step takes 30 min and the LysoTracker staining 10 min, so it can be performed relatively rapidly. As data are typically collected on a minimum of 10,000 live cells, the flow cytometry provides robust quantitative data. Another advantage is that the flow cytometry data contain physical parameters about the blood cells in the sample. You can therefore also see if there is a major change in other hematopoietic cell types over time, which may be useful information for longitudinal monitoring of disease progression/response to therapy. The assay could be adapted to a 96-well fluorescent plate reader format as long as cell loading can be controlled. However, fluorescent plate readers lack the very large dynamic range (5-decade log scale) offered by flow cytometry, which confers the high level of sensitivity needed, so will not be discussed further in this chapter. The downside to this approach is the need to work with live cells, so again blood samples have to be analyzed within a certain window of time (see below). However, one major advantage of the use of this live cell-staining approach is that dead cells do not maintain LE/Lys pH, so do not stain with LysoTracker resulting in only viable cells being stained. The assay can be applied to vertebrate models of LSDs and patient blood samples, and so methods for mouse and human are provided to illustrate this. If studying
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large animal models of LSDs, make sure there is a good pan-B-cell antibody available for the species in question, as this is a pre-requisite for the assay.
4. METHODS An overview of the protocol is shown schematically in Figure 1.
4.1 PREPARATION OF HUMAN BLOOD 4.1.1 Materials and reagents Blood collected into ethylenediaminetetraacetic acid (EDTA) tubes (“purple cap” Becton Dickinson (BD) vacutainer blood collection tubes) 15-mL centrifuge tubes 1.5-mL centrifuge tubes Histopaque-1077 (SigmaeAldrich) Disposable plastic Pasteur pipettes Dulbecco’s phosphate buffered saline (D-PBS) (SigmaeAldrich)
FIGURE 1 Schematic representation of the assay. The assay starts with blood collection, through to mononuclear cell separation then LysoTracker/B-cell staining followed by flow cytometric analysis. There is also the optional step of isolating B-cells using magnetic beads, which can then be analyzed by microscopy or stored frozen for future biochemical analysis.
4. Methods
FIGURE 2 Schematic representation of a blood tube after centrifugation.
Hemocytometer 0.5% Trypan blue
4.1.2 Protocol • • • • • • • • • • •
Load 3 mL whole blood onto 3 mL Histopaque-1077 (warmed to room temperature (RT)) in a 15-mL tube using a disposable plastic Pasteur pipette. Spin the tubes for 30 min at 400 x g at RT (20 C). The tubes should come out of the centrifuge resembling Figure 2. Remove the plasma using a pipette. Transfer the white blood cells and platelets fraction into a 15-mL tube, add 10 mL D-PBS, and gently mix the tube. Spin the tubes for 10 min at 250 x g. Remove the supernatant (the upper layer is still opaque, due to the platelets). To wash out all the Histopaque, add 13 mL D-PBS to the pellet, resuspend the pellet, and spin once more for 10 min at 250 x g. Remove the supernatant and resuspend the cell pellets in 0.5 mL D-PBS. Take 10 mL of the mononuclear cells, add 80 mL D-PBS and 10 mL 0.5% trypan blue, mix and use this mixture to count the cells on a hemocytometer. Use one million cells in duplicate for the CD19/LysoTracker assay.
4.2 PREPARATION OF CELLS FROM MOUSE SPLEEN OR WHOLE BLOOD There are many authentic mouse models of LSDs (Hemsley & Hopwood, 2010) that can also be studied using this assay (te Vruchte et al., 2014), and both blood and spleen are good sources of B-cells.
4.2.1 Materials and reagents Twin frosted microscope slides Petri dishes
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D-PBS (SigmaeAldrich) 15-mL centrifuge tubes 50-mL centrifuge tubes Red blood cell lysis buffer (0.14 M NH4Cl, 0.017 M Tris-Base pH7.2) Single cell sieves (70 mM) Hemocytometer 0.5% Trypan blue.
4.2.2 Protocol 4.2.2.1 Isolation of mononuclear cells from mouse spleen •
•
• • •
• •
Make a suspension of the mouse spleen (freshly collected on the day and kept on ice) by cutting the spleen into pieces with scissors, then massaging the pieces between two frosted ends of two microscope slides above a petri dish with 10 mL D-PBS, now and then dipping the slides in the D-PBS. Transfer the cell suspension to a 15-mL tube and spin the cells for 5 min at 750 x g. Resuspend the cells in 5 mL red blood cell lysis buffer and incubate for 10 min at RT (lysis only works at RT, so make sure the lysis buffer is taken from the fridge ahead of time). Sieve this through a single cell sieve into a 50-mL tube with 10 mL D-PBS. Spin down the cells 5 min at 750 x g (if the pellet is still red, then repeat the lysis) and resuspend the cells in 1 mL D-PBS. *Optional step: If for any reason the FACS analysis cannot be conducted on the same day, the cells can be resuspended in 1 mL FCS þ 10% DMSO, frozen in cryotubes (as you would to freeze down cell lines) at 80 C overnight and then transferred to liquid nitrogen. Typically, these cells are thawed and analyzed within 30 days. Take 10 mL of the cells, add 80 mL D-PBS and 10 mL 0.5% trypan blue, mix and count the cells on a hemocytometer. Use one million cells in duplicate for the CD19/LysoTracker assay. * NB. When the single cells have been frozen in FCS þ 10% dimethyl sulfoxide (DMSO): • • • • •
Remove the cells from the liquid nitrogen. Rapidly warm to 37 C in a water bath. Wash the cells three times with D-PBS þ CaCl2 þ MgCl (Sigma). Count the cells. Take one million cells for staining
4.2.2.2 Isolation of mononuclear cells from mouse blood • • •
Collect the blood from the mouse (300e1000 mL) and put this into a 15-mL tube with 5 mL D-PBS þ 5 mM EDTA. Spin the cells for 5 min at 750 x g. Resuspend the cells in 5 mL red blood cell lysis buffer and incubate for 10 min at RT.
4. Methods
• • • •
Make the volume up to 14 mL with D-PBS. Spin down the cells, repeat the lysis step once more, spin and resuspend the cells in 0.5 mL D-PBS. Take 10 mL of the cells, add 80 mL D-PBS and 10 mL 0.5% trypan blue, mix and use this mixture to count the cells on a hemocytometer. Use one million cells for the CD19/LysoTracker assay. Mouse blood cells can also be cryopreserved for later analysis (see above).
4.3 B-LYMPHOCYTE STAINING 4.3.1 Materials and reagents for human cells PE-conjugated mouse anti-human CD19 antibody (clone LT19; Abcam) D-PBS (SigmaeAldrich) 15-mL centrifuge tubes 10% BSA FACS buffer (0.1% BSA, 0.02 M NaN3 in 1 PBS) 200 nM LysoTrackerÒ-green DND-26 (Invitrogen)
4.3.2 Protocol for human cells • • • • • • • • • •
Incubate one million mononuclear cells in a volume of 100 mL with 15 mL PE-conjugated mouse antihuman CD19 antibody in the dark on ice for 30 min. After 30 min, add 100 mL 10% BSA. Spin the cells for 5 min at 800 x g. Remove the supernatant. Resuspend the cells in 1 mL of 200 nM LysoTrackerÒ-green DND-26. Leave this in the dark at RT for 10 min. Spin the cells for 5 min at 800 x g. Remove the supernatant. Resuspend the cells in 500 mL FACS buffer and transfer the cells into FACS tubes. Put the cells on ice and analyze them on the FACS as soon as possible in a standardized way every time you run the assay (see below).
4.3.3 Materials and reagents for mouse cells Fc-block (0.5 mg/mL purified anti-mouse CD16/CD32; BD Pharmingen) CD19-PE (0.2 mg/mL PE anti-mouse CD19; BD Pharmingen) D-PBS (SigmaeAldrich) 15-mL centrifuge tubes 10% BSA FACS buffer (0.1% BSA, 0.02 M NaN3 in 1 PBS) 200 nM LysoTrackerÒ-green DND-26 (Invitrogen) FACS tubes
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4.3.4 Protocol for mouse cells • • • • • • • • • • •
Incubate one million cells in a volume of 50 mL with 2 mL Fc-block on ice for 5 min. Add 5 mL CD19-PE and incubate for 30 min in the dark on ice. Stop the incubation by adding 100 mL 10% BSA. Spin the cells for 5 min at 800 x g. Remove the supernatant. Resuspend the cells in 1 mL of 200 nM LysoTrackerÒ-green DND-26. Leave this in the dark at RT for 10 min. Spin the cells for 5 min at 800 x g. Remove the supernatant. Resuspend the cells in 500 mL FACS buffer and transfer the cells into FACS tubes. Put the cells on ice and analyze them on the FACS as soon as possible in a standardized way every time you run the assay (see below).
4.4 FLOW CYTOMETRY, CALIBRATION, ACQUISITION, AND ANALYSIS We use a BD Biosciences FACSCanto II for all our flow cytometric analysis, but other flow cytometer instruments can be used. Representative images showing the Fluorescence-activated cell sorting (FACS) analysis are shown in Figure 3. It is best however to use the same machine for all analysis if you need to compare the results over time as absolute values rather than as fold-changes (see discussion below).
4.4.1 Materials and reagents BD Biosciences FACSCanto II Cytometer Setup and Tracking beads (BD) Anti-mouse CompBeads (Anti-Mouse Ig, k/Negative Control (FBS) Compensation Particles Set, BD) 8-peak Rainbow calibration beads (SPHEROÔ Rainbow Calibration Particles, BD)
4.4.2 Protocol • • •
• •
Calibrate the cytometer using Cytometer Setup and Tracking beads (BD). Compensation is performed using cells stained with LysoTracker and anti-mouse CompBeads (BD) stained with PE antibody using FACSDiva software (BD). Samples are acquired with gating on singlet cells (FSC-H vs FSC-A) and CD19þ events. In total, 50,000 singlet events and 10,000 singlet gate CD19þ events are collected. The mean fluorescence of the CD19þ events is calculated using FACSDiva software (BD). The molecules of equivalent fluorescence (MEFL) is calculated using 8-peak Rainbow calibration beads (BD), using the fluorescein equivalent values provided by the manufacturer.
This shows singlet gating, Forward Scatter (FSC) versus Side Scatter (SSC) for human blood, the CD19 population identified with anti-CD19 antibody staining (red) and LysoTracker staining in green, and histogram of molecules of equivalent fluorescence (MEFL) of LysoTracker on B-cell population. (See color plate)
4. Methods
FIGURE 3 Representative images of flow cytometry setup windows.
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4.5 ISOLATION OF B-CELLS FOR BIOCHEMICAL ASSAYS OR MICROSCOPY When B-cells are needed for biochemical assays or for microscopy, they can be isolated from the mononuclear cell fraction isolated (as described above), using magnetic beads. We use magnetic cell separation (MACS) beads from Miltenyi Biotec for this purpose. We have used Dynabeads Mouse pan B (Life Technologies) in the past and found that these beads were fluorescent under a microscope, but are fine if biochemical analysis is to be performed, not microscopy.
4.5.1 Materials and reagents for human cells MACS buffer (D-PBS supplemented with 0.5% BSA and 2 mM EDTA) MACS Human CD19 MicroBeads (Miltenyi Biotec) 15-mL centrifuge tubes Tube roller bank (Miltenyi Biotec) MS column (Miltenyi Biotec) MACS separator (Miltenyi Biotec)
4.5.2 Protocol for human cells • • •
• • • • •
• •
Resuspend the mononuclear cells in 80 mL of MACS buffer per 10E7 total cells. Add 20 mL of MACS Human CD19 MicroBeads per 10E7 total cells, mix well, and incubate for 15 min on a tube roller bank at 4 C. Wash the cells by adding 10e20 x the labeling volume of buffer, centrifuge at 300 x g for 10 min, remove the supernatant completely, and resuspend the cell pellet in 500 mL MACS buffer. Place an MS column in the magnetic field of a MACS separator. Wash the column with 500 ml MACS buffer. Apply the cell suspension to the column. Rinse three times with 500 mL MACS buffer. Remove the column from the separator, place the column on a 15-mL tube, pipette 1 mL of MACS buffer onto the column, and flush out the positive cells using the plunger supplied with the column. Count the B-cells. Spin down the cells, remove the supernatant and wash them once with 1 mL D-PBS, remove the supernatant again and freeze the pellet at 80 C for biochemical analysis later or use the cells straightaway for staining for microscopy.
4.5.3 Materials and reagents for mouse cells MACS buffer (D-PBS supplemented with 0.5% BSA and 2 mM EDTA) MACS Mouse CD19 MicroBeads (Miltenyi Biotec) 15-mL centrifuge tubes Tube roller bank (Miltenyi Biotec) MS column (Miltenyi Biotec) MACS separator (Miltenyi Biotec)
4. Methods
4.5.4 Protocol for mouse cells • • •
• • • • •
•
Resuspend the mononuclear cells in 90 mL of MACS buffer per 10E7 total cells. Add 10 mL of MACS Mouse CD19 MicroBeads per 10E7 total cells, mix well, and incubate for 15 min on a tube roller bank at 4 C. Wash the cells by adding 10e20x the labeling volume of buffer, centrifuge at 300 x g for 10 min, remove the supernatant completely, and resuspend the cell pellet in 500 mL MACS buffer. Place an MS column in the magnetic field of a MACS separator. Wash the column with 500 mL MACS buffer. Apply the cell suspension to the column. Rinse three times with 500 mL MACS buffer. Remove the column from the separator, place the column on a 15-mL tube, pipette 1 mL of MACS buffer onto the column, and flush out the positive cells using the plunger supplied with the column. Spin down the cells, remove the supernatant and wash them once with 1 mL D-PBS, remove the supernatant again and freeze the pellet at 80 C for biochemical analysis later or use the cells straightaway for staining for microscopy.
4.6 INFLUENCE OF PATIENT BLOOD SHIPPING TIMES One of the practical considerations before implementing this method in your laboratory is to consider the effect of blood storage/shipping times on assay performance. Several practical scenarios can be envisaged. In a clinical setting, it may be possible to draw and analyze patient blood the same day in which case the situation is straightforward. However, even in this scenario it may not be possible to perform the assay on the same day depending on clinic visit times for the patient and so the question is: How should the sample be stored and does storage affect the results? This is even more of an issue in a research setting where samples will be shipped to the research laboratory for analysis from a clinic that may be in another country. This involves international shipments and considerable lag times between drawing the blood and their arrival in the research laboratory for analysis. The two key variables to consider are the blood sample storage/shipping temperature and its influence on assay performance and also how long a sample can be in transit and still be suitable for assay. We recently completed a longitudinal study spanning many years on Niemann-Pick type C patient blood samples and had to deal with these practical issues (te Vruchte et al. 2014). What we will do is share our experiences with you, so they can inform your choices for your own studies/own situation and show you some experimental data on how temperature and time affect the analysis. The simple take-home message is you must adopt one protocol and stick to it rigidly to allow accurate comparisons to be made between samples over time.
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4.7 INFLUENCE OF BLOOD STORAGE TEMPERATURE ON LYSOTRACKER STAINING Blood samples collected into EDTA vacutainer tubes can be stored at RT or can be refrigerated at 4 C. As our study (te Vruchte et al. 2014) required shipping of samples from Germany, the UK, and the USA, we defaulted to RT, as it was simpler for the clinics to deal with. The downside was that this was prone to changes in ambient temperature at airports depending on the time of year but generally speaking was in the 20e25 C range. Here we have run a small experiment to show you how temperature affects the assay and why you need to standardize this parameter. We drew blood from healthy volunteers and compared LysoTracker staining values (MEFL) in their B-cells when we stored the blood at RT or 4 C for up to 5 days. The samples were assayed daily as described above. The data are summarized in Figure 4. The samples maintained at 4 C were less affected by storage than RT samples, with the major reduction in MEFL signal occurring over the first 24 h. Interestingly, B-cells surviving out to 5 days albeit low in number gave very robust MEFL values comparable to B-cells assayed on day 1e2 (we since repeated this and made the same observation, not shown). As LysoTracker is a live cell stain, there is no need to worry about dead cells in this assay. The take-home message is as long as one temperature and one analysis time (days post-blood draw) are chosen, data over time and between individuals can be reliably compared. Fold change over time rather than
FIGURE 4 Effect of storing blood samples at 4 C or room temperature for 5 days. Samples were collected from five healthy volunteers and molecules of equivalent fluorescence (MEFL) of B-cells assayed by FACS on a daily basis for 5 days.
4. Methods
absolute values can also be used to compare data between centers. Flow cytometers also vary, so absolute comparisons will be difficult to make between instruments. Again, fold change relative to control samples can be used reliably.
4.8 INFLUENCE OF DELAYS IN ANALYSIS OF SAMPLES POST-LYSOTRACKER STAINING Another potential variable is the lag time between staining with LysoTracker and actually analyzing the samples on the flow cytometer. We evaluated this variable (Figure 5), and rapid analysis times gave the strongest signals. It is therefore important to standardize this carefully in any protocol you establish. Long delays give reduced sensitivity (low MEFL values), so analyzing within the first 30 min is recommended and should be kept constant from assay to assay.
4.8.1 Some economic considerations relating to B-cell staining The assay we describe is low cost (assuming you have a flow cytometer already in place) except for the LysoTracker and the anti-CD19 reagents. We initially followed manufacturer’s instructions for the anti-CD19 staining, but as you can see in Figure 6 we were able to titrate the antibody considerably and still achieve robust delineation of the CD19-positive population. So titrating the antibody you purchase can save
FIGURE 5 Effect of delaying analysis after LysoTracker staining. Analysis of samples within 30 min gave optimal signal intensity, which then declined up to 2 h post-staining. This illustrates the need to rapidly analyze samples after live cell staining with LysoTracker and the need to standardize the protocol, so this does not vary between experiments.
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FIGURE 6 Titration of anti-CD19 antibody. As can be seen, considerable titration of anti-CD19 was possible, well below the manufacturers recommended antibody concentration of 20 mL.
you money. The only requirement is that you use enough antibody to be able to identify the B-cells as a discreet population.
5. SUMMARY Relative lysosomal volume can be readily measured in circulating B-cells in mouse and human blood samples for studying LSDs. As long as you adopt a rigidly standardized protocol in relation to day of analysis post-blood draw, temperature the blood is stored/shipped at, and time from addition of LysoTracker to analysis on the cytometer, the assay is robust. We suggest that when comparing data between centers, where different cytometers/protocol times etc., are being used, a fold change from baseline or age-matched controls is the most pragmatic solution for comparing data rather than absolute values.
ACKNOWLEDGMENTS DtV and FMP were supported by Action Medical Research, Niemann-Pick UK and an unrestricted grant from Actelion. KLW is supported by a Stratified Medicine Grant (Gaucherite) from the MRC. FMP is a Royal Society Wolfson Research Merit Award holder. We thank the volunteers who donated blood for this study.
References
REFERENCES Beutler, E. (2006). Lysosomal storage diseases: natural history and ethical and economic aspects. Molecular Genetics and Metabolism, 88, 208e215. Bright, N. A., Gratian, M. J., & Luzio, J. P. (2005). Endocytic delivery to lysosomes mediated by concurrent fusion and kissing events in living cells. Current Biology, 15, 360e365. Calcraft, P. J., Ruas, M., Pan, Z., Cheng, X., Arredouani, A., Hao, X., et al. (2009). NAADP mobilizes calcium from acidic organelles through two-pore channels. Nature, 459, 596e600. Churchill, G. C., Okada, Y., Thomas, J. M., Genazzani, A. A., Patel, S., & Galione, A. (2002). NAADP mobilizes Ca(2þ) from reserve granules, lysosome-related organelles, in sea urchin eggs. Cell, 111, 703e708. Gadola, S. D., Silk, J. D., Jeans, A., Illarionov, P. A., Salio, M., Besra, G. S., et al. (2006). Impaired selection of invariant natural killer T cells in diverse mouse models of glycosphingolipid lysosomal storage diseases. Journal of Experimental Medicine, 203, 2293e2303. Hemsley, K. M., & Hopwood, J. J. (2010). Lessons learnt from animal models: pathophysiology of neuropathic lysosomal storage disorders. Journal of Inherited Metabolic Disease, 33, 363e371. Lachmann, R. H., te Vruchte, D., Lloyd-Evans, E., Reinkensmeier, G., Sillence, D. J., Fernandez-Guillen, L., et al. (2004). Treatment with miglustat reverses the lipid-trafficking defect in Niemann-Pick disease type C. Neurobiology of Disease, 16, 654e658. Lloyd-Evans, E., Morgan, A. J., He, X., Smith, D. A., Elliot-Smith, E., Sillence, D. J., et al. (2008). Niemann-Pick disease type C1 is a sphingosine storage disease that causes deregulation of lysosomal calcium. Nature Medicine, 14, 1247e1255. Luzio, J. P., Pryor, P. R., & Bright, N. A. (2007). Lysosomes: fusion and function. Nature Reviews Molecular Cell Biology, 8, 622e632. Platt, F. M. (2014). Sphingolipid lysosomal storage disorders. Nature, 510, 68e75. Platt, F. M., Boland, B., & van der Spoel, A. C. (2012). The cell biology of disease: lysosomal storage disorders: the cellular impact of lysosomal dysfunction. Journal of Cell Biology, 199, 723e734. Speak, A. O., Te Vruchte, D., Davis, L. C., Morgan, A. J., Smith, D. A., Yanjanin, N. M., et al. (2014). Altered distribution and function of natural killer cells in murine and human Niemann-Pick disease type C1. Blood, 123, 51e60. te Vruchte, D., Speak, A. O., Wallom, K. L., Al Eisa, N., Smith, D. A., Hendriksz, C. J., et al. (2014). Relative acidic compartment volume as a lysosomal storage disorder-associated biomarker. Journal of Clinical Investigation, 124, 1320e1328. Vitner, E. B., Platt, F. M., & Futerman, A. H. (2010). Common and uncommon pathogenic cascades in lysosomal storage diseases. Journal of Biological Chemistry, 285, 20423e20427. Winchester, B., Vellodi, A., & Young, E. (2000). The molecular basis of lysosomal storage diseases and their treatment. Biochemical Society Transactions, 28, 150e154. Wraith, J. E. (2002). Lysosomal disorders. Seminars in Neonatology, 7, 75e83. Zoncu, R., Bar-Peled, L., Efeyan, A., Wang, S., Sancak, Y., & Sabatini, D. M. (2011). mTORC1 senses lysosomal amino acids through an inside-out mechanism that requires the vacuolar H-ATPase. Science, 334, 678e683.
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Measuring relative lysosomal volume for monitoring lysosomal storage diseases
16
Danielle te Vruchte1, Kerri L. Wallom, Frances M. Platt1 1
Department of Pharmacology, University of Oxford, Oxford, UK
Corresponding authors: E-mail: [email protected]; [email protected]
CHAPTER OUTLINE 1. 2. 3. 4.
Measuring Relative Lysosomal Volume as an Index of Lysosomal Storage .............. 332 In Which Circulating Cell Type Should Relative Lysosomal Volume Be Measured?.. 333 How to Measure Relative Lysosomal Volume in Blood Cells .................................. 334 Methods ............................................................................................................ 336 4.1 Preparation of Human Blood ................................................................ 336 4.1.1 Materials and reagents ..................................................................... 336 4.1.2 Protocol ........................................................................................... 337 4.2 Preparation of Cells from Mouse Spleen or Whole Blood ......................... 337 4.2.1 Materials and reagents ..................................................................... 337 4.2.2 Protocol ........................................................................................... 338 4.3 B-Lymphocyte Staining........................................................................ 339 4.3.1 Materials and reagents for human cells ............................................. 339 4.3.2 Protocol for human cells ................................................................... 339 4.3.3 Materials and reagents for mouse cells ............................................. 339 4.3.4 Protocol for mouse cells.................................................................... 340 4.4 Flow Cytometry, Calibration, Acquisition, and Analysis ........................... 340 4.4.1 Materials and reagents ..................................................................... 340 4.4.2 Protocol ........................................................................................... 340 4.5 Isolation of B-Cells for Biochemical Assays or Microscopy....................... 342 4.5.1 Materials and reagents for human cells ............................................. 342 4.5.2 Protocol for human cells ................................................................... 342 4.5.3 Materials and reagents for mouse cells ............................................. 342 4.5.4 Protocol for mouse cells.................................................................... 343 4.6 Influence of Patient Blood Shipping Times ............................................ 343 4.7 Influence of Blood Storage Temperature on LysoTracker Staining ............ 344
Methods in Cell Biology, Volume 126, ISSN 0091-679X, http://dx.doi.org/10.1016/bs.mcb.2014.10.027 © 2015 Elsevier Inc. All rights reserved.
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4.8 Influence of Delays in Analysis of Samples Post-LysoTracker Staining...... 345 4.8.1 Some economic considerations relating to B-cell staining .................. 345 5. Summary ........................................................................................................... 346 Acknowledgments ................................................................................................... 346 References ............................................................................................................. 347
Abstract Biomarkers are important tools in medicine, which can be used for monitoring disease progression and response to therapy. One of the main problems in rare lysosomal storage diseases is that there are over 70 different diseases, all with different biochemical storage profiles. Developing biochemical biomarkers therefore requires an individual assay per disease/subgroup of diseases. An alternative approach is to develop an assay that is independent of the specific macromolecules stored. This chapter discusses an assay that may serve as a universal biomarker for these diseases and measures the expansion of the late endosomal/lysosomal system. We have developed an assay that takes advantage of a commercially available late endosomal/lysosomal probe, LysoTracker, which becomes trapped in the acidic compartment of cells and emits a fluorescent signal that can be detected using flow cytometry. In this chapter, we detail the methodology behind this assay and discuss the factors that need to be considered when establishing this assay in clinical and research settings.
1. MEASURING RELATIVE LYSOSOMAL VOLUME AS AN INDEX OF LYSOSOMAL STORAGE The late endocytic system comprises two main acidic organelles, namely late endosomes (LE) and lysosomes (Lys). Their volume is tightly regulated in a cell-typespecific manner. Resting cells tend to have a lower LE/Lys volume compared to highly active cells (e.g., macrophages). The function of LE/Lys is the catabolism and recycling of cellular macromolecules (Luzio, Pryor, & Bright, 2007). However, in addition they serve as nutrient sensors (Zoncu et al., 2011) and are signaling organelles in part through their role as regulated calcium stores (Churchill et al., 2002; Calcraft et al., 2009). Irrespective of the cell type, if there is an inborn error of metabolism that affects lysosomal function, substrates and cargos within LE/Lys build up leading to “storage” and disease (Platt, Boland, & van der Spoel, 2012). Typically, these inherited diseases are caused by mutations in genes encoding lysosomal enzymes (acid hydrolases), but defects in non-enzymatic lysosomal proteins (soluble or membrane proteins) can also lead to lysosomal storage (Winchester, Vellodi, & Young, 2000; Wraith, 2002; Beutler, 2006). The accumulation of non-degraded metabolites in the LE/Lys system then triggers an increase in LE/Lys volume, providing a potentially universal cellular biomarker for these diseases (Vitner, Platt, & Futerman, 2010). The lysosomal storage diseases (LSDs) comprise >70 individually rare diseases, but are collectively much more common and affect approximately 1:5000 live births
2. Measuring relative lysosomal volume as an index of storage
(Vitner et al. 2010; Platt et al. 2012). The technical advantage of being able to measure relative changes in acidic compartment volume is that it allows storage levels to be measured in patient cells without the need to measure disease-specific biochemical metabolites. Therapies for these diseases aim to reduce LE/Lys storage; therefore, measuring relative lysosomal volume can also potentially report on response to treatment. The current approved therapies for LSDs are enzyme replacement therapy, bone marrow transplantation, and substrate reduction therapy using small molecules (Platt et al. 2012; Platt, 2014). However, other therapeutics are undergoing clinical trials at the current time (gene therapy, chaperone therapies, etc.) and have recently been reviewed elsewhere (Platt, 2014). The main practical applications of measuring LE/Lys volume as a potentially universal cellular biomarker for LSDs can be summarized as follows: 1. Suspicion of an LSD: Based on clinical presentation, this can be confirmed at the cellular level in a blood sample or in skin fibroblasts by measuring LE/Lys volume. This can be performed prior to commissioning more costly diseasespecific diagnostic tests, typically enzyme assays and mutation analysis. 2. Following diagnosis of an LSD, peripheral blood cells from the patient can be measured over time, allowing longitudinal progression of storage to be measured quantitatively. 3. In clinical trials, the use of this biomarker may aid stratification of patients for recruitment to trials and also for monitoring individual patient responses to a clinical intervention. 4. In LSDs with approved therapies, relative lysosomal volume can also be used as a monitoring tool to study long-term stability and detect any signs of a failure in treatment efficacy over time. 5. As many LSDs may benefit from combination therapies, in the future this could also be a useful biomarker for determining whether the addition of a new therapeutic agent significantly impacts levels of storage. As blood is a minimally invasive source of patient cells, it is an ideal starting point for such an assay. However, blood contains a great variety of cells of hematopoietic origin, and a very important question is therefore: Which circulating nucleated cell type should be monitored? A second question is: How can relative lysosomal volume be measured in blood cells? We will discuss both of these questions in more detail before discussing the detailed methods.
2. IN WHICH CIRCULATING CELL TYPE SHOULD RELATIVE LYSOSOMAL VOLUME BE MEASURED? What proved to be the most important factor in the reliability and reproducibility of the assay we have developed was the choice of cell type to analyze (Lachmann et al., 2004; te Vruchte et al., 2014). All mononuclear cells have LE/Lys compartments, so are all candidates to be measured? Readers may be thinking why not simply profile
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all mononuclear cells in human blood and take an average or get data on multiple cell types/populations. The reason this is not a good approach is that the hematopoietic system varies significantly with age (a newborn does not have the same circulating blood cell ratios as an infant or an adult) and most significantly these cells change with disease progression in LSDs (Gadola et al., 2006; Speak et al., 2014). Therefore, a specific cell type needs selecting for LE/Lys volume determination to ensure reliable longitudinal analysis in the same individual and permit valid comparisons to be made between individuals. We originally focused on cells with the largest LE/Lys volume (myeloid cells), but they can only be accurately identified using multiple cell surface markers. Very significantly, they change in response to infection to a variety of different activation states, requiring even more markers to be analyzed in order to identify them accurately. It is therefore difficult to be sure that precisely the same cell type is being monitored over time making them a poor choice for this cellular assay. In the end, we focused our attention on small circulating resting B-cells and they indeed proved ideal for our purposes (Lachmann et al. 2004; te Vruchte et al. 2014). Small resting B-cells are antigen naı¨ve and have a very low cytoplasmic-to-nuclear volume. Their acidic compartment volume is therefore relatively low compared with myeloid cells, but they have the major advantage that they all express the pan-B-cell marker CD19. This means that they can be reliably identified with a single antibody without the need to sort or purify the cells, and they are compatible with longitudinal analysis, as they are a homogeneous population that crucially does not change in response to infection.
3. HOW TO MEASURE RELATIVE LYSOSOMAL VOLUME IN BLOOD CELLS There are at least three main ways that relative lysosomal volume could be measured in mononuclear cells: (1) by measuring a lysosomal protein by western blot or enzymelinked immunosorbent assay (ELISA) as a reporter on relative compartment size/ volume, (2) by measuring a fluorescent probe that is endocytosed and “chased” into the late endocytic system, or (3) by using a cell-permeant fluorescent probe that is rapidly trapped in acidic organelles and can be measured to quantify relative volume of LE/Lys. All of these approaches are equally valid but have a number of pros and cons from a practical perspective. Measuring lysosomal proteins by western blotting assumes that the level of the lysosomal protein of interest scales with LE/Lys volume in a given LSD. This method requires the isolation of the cell type to be measured free of contaminating cells; the proteins then need extracting with detergent containing buffer, running on an sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) gel, transferring to a suitable membrane, and probing with an appropriate antibody to detect the lysosomal protein of interest. This is a time-consuming exercise and low throughput. It is also relatively expensive due to the need to run gels, to blot
3. How to measure relative lysosomal volume in blood cells
and probe the blot with a reasonable volume of antibody. ELISAs have a similar limitation in that the cells need isolating to high purity, but in this case there is no need to run gels, as the lysosomal protein is simply measured in an ELISA plate. Of the two methods, ELISAs are higher throughput, but the cell isolation step is the big drawback. Even a small contamination with myeloid cells can distort the lysosomal marker levels in a B-cell preparation very significantly. However, the advantage of these approaches is that they can be performed on frozen material, as long as the cell purification is performed on fresh cells. The alternative is to pulse the cells in question with a fluorescent reporter that is “chased” into LE/Lys. Fluorescent dextran, for example, could be used for this purpose (Bright, Gratian, & Luzio, 2005; Lloyd-Evans et al., 2008). However, pulsechase times need to be optimized and the chase time significantly extends the assay time. Again, making sure a single homogeneous cell population is studied is crucial, as the chase times will be cell-type dependent. This requires live cell analysis by microscopy, so they must be analyzed before viability is compromised. The analysis is tedious and not particularly quantitative. The method we have adopted and optimized uses a cell-permeant dye, such as LysoTracker, which is a weak amine. At acidic pH, it is protonated and becomes trapped in LE/Lys. The relative fluorescence of the signal is then proportional to the relative volume of the LE/Lys compartment (te Vruchte et al., 2014). There are several advantages of this method. Firstly, it can be coupled with very sensitive analytical flow cytometry that has a large dynamic range. B-cells are simply identified in the circulating mononuclear cell populations with an anti-CD19 antibody (e.g., red conjugate). The cells are then co-labeled with LysoTracker (e.g., LysoTracker green) and the B-cell population analyzed without the need to sort or purify B-cells using other methods. This greatly speeds up the assay. The second advantage is that the antibody-staining step takes 30 min and the LysoTracker staining 10 min, so it can be performed relatively rapidly. As data are typically collected on a minimum of 10,000 live cells, the flow cytometry provides robust quantitative data. Another advantage is that the flow cytometry data contain physical parameters about the blood cells in the sample. You can therefore also see if there is a major change in other hematopoietic cell types over time, which may be useful information for longitudinal monitoring of disease progression/response to therapy. The assay could be adapted to a 96-well fluorescent plate reader format as long as cell loading can be controlled. However, fluorescent plate readers lack the very large dynamic range (5-decade log scale) offered by flow cytometry, which confers the high level of sensitivity needed, so will not be discussed further in this chapter. The downside to this approach is the need to work with live cells, so again blood samples have to be analyzed within a certain window of time (see below). However, one major advantage of the use of this live cell-staining approach is that dead cells do not maintain LE/Lys pH, so do not stain with LysoTracker resulting in only viable cells being stained. The assay can be applied to vertebrate models of LSDs and patient blood samples, and so methods for mouse and human are provided to illustrate this. If studying
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large animal models of LSDs, make sure there is a good pan-B-cell antibody available for the species in question, as this is a pre-requisite for the assay.
4. METHODS An overview of the protocol is shown schematically in Figure 1.
4.1 PREPARATION OF HUMAN BLOOD 4.1.1 Materials and reagents Blood collected into ethylenediaminetetraacetic acid (EDTA) tubes (“purple cap” Becton Dickinson (BD) vacutainer blood collection tubes) 15-mL centrifuge tubes 1.5-mL centrifuge tubes Histopaque-1077 (SigmaeAldrich) Disposable plastic Pasteur pipettes Dulbecco’s phosphate buffered saline (D-PBS) (SigmaeAldrich)
FIGURE 1 Schematic representation of the assay. The assay starts with blood collection, through to mononuclear cell separation then LysoTracker/B-cell staining followed by flow cytometric analysis. There is also the optional step of isolating B-cells using magnetic beads, which can then be analyzed by microscopy or stored frozen for future biochemical analysis.
4. Methods
FIGURE 2 Schematic representation of a blood tube after centrifugation.
Hemocytometer 0.5% Trypan blue
4.1.2 Protocol • • • • • • • • • • •
Load 3 mL whole blood onto 3 mL Histopaque-1077 (warmed to room temperature (RT)) in a 15-mL tube using a disposable plastic Pasteur pipette. Spin the tubes for 30 min at 400 x g at RT (20 C). The tubes should come out of the centrifuge resembling Figure 2. Remove the plasma using a pipette. Transfer the white blood cells and platelets fraction into a 15-mL tube, add 10 mL D-PBS, and gently mix the tube. Spin the tubes for 10 min at 250 x g. Remove the supernatant (the upper layer is still opaque, due to the platelets). To wash out all the Histopaque, add 13 mL D-PBS to the pellet, resuspend the pellet, and spin once more for 10 min at 250 x g. Remove the supernatant and resuspend the cell pellets in 0.5 mL D-PBS. Take 10 mL of the mononuclear cells, add 80 mL D-PBS and 10 mL 0.5% trypan blue, mix and use this mixture to count the cells on a hemocytometer. Use one million cells in duplicate for the CD19/LysoTracker assay.
4.2 PREPARATION OF CELLS FROM MOUSE SPLEEN OR WHOLE BLOOD There are many authentic mouse models of LSDs (Hemsley & Hopwood, 2010) that can also be studied using this assay (te Vruchte et al., 2014), and both blood and spleen are good sources of B-cells.
4.2.1 Materials and reagents Twin frosted microscope slides Petri dishes
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D-PBS (SigmaeAldrich) 15-mL centrifuge tubes 50-mL centrifuge tubes Red blood cell lysis buffer (0.14 M NH4Cl, 0.017 M Tris-Base pH7.2) Single cell sieves (70 mM) Hemocytometer 0.5% Trypan blue.
4.2.2 Protocol 4.2.2.1 Isolation of mononuclear cells from mouse spleen •
•
• • •
• •
Make a suspension of the mouse spleen (freshly collected on the day and kept on ice) by cutting the spleen into pieces with scissors, then massaging the pieces between two frosted ends of two microscope slides above a petri dish with 10 mL D-PBS, now and then dipping the slides in the D-PBS. Transfer the cell suspension to a 15-mL tube and spin the cells for 5 min at 750 x g. Resuspend the cells in 5 mL red blood cell lysis buffer and incubate for 10 min at RT (lysis only works at RT, so make sure the lysis buffer is taken from the fridge ahead of time). Sieve this through a single cell sieve into a 50-mL tube with 10 mL D-PBS. Spin down the cells 5 min at 750 x g (if the pellet is still red, then repeat the lysis) and resuspend the cells in 1 mL D-PBS. *Optional step: If for any reason the FACS analysis cannot be conducted on the same day, the cells can be resuspended in 1 mL FCS þ 10% DMSO, frozen in cryotubes (as you would to freeze down cell lines) at 80 C overnight and then transferred to liquid nitrogen. Typically, these cells are thawed and analyzed within 30 days. Take 10 mL of the cells, add 80 mL D-PBS and 10 mL 0.5% trypan blue, mix and count the cells on a hemocytometer. Use one million cells in duplicate for the CD19/LysoTracker assay. * NB. When the single cells have been frozen in FCS þ 10% dimethyl sulfoxide (DMSO): • • • • •
Remove the cells from the liquid nitrogen. Rapidly warm to 37 C in a water bath. Wash the cells three times with D-PBS þ CaCl2 þ MgCl (Sigma). Count the cells. Take one million cells for staining
4.2.2.2 Isolation of mononuclear cells from mouse blood • • •
Collect the blood from the mouse (300e1000 mL) and put this into a 15-mL tube with 5 mL D-PBS þ 5 mM EDTA. Spin the cells for 5 min at 750 x g. Resuspend the cells in 5 mL red blood cell lysis buffer and incubate for 10 min at RT.
4. Methods
• • • •
Make the volume up to 14 mL with D-PBS. Spin down the cells, repeat the lysis step once more, spin and resuspend the cells in 0.5 mL D-PBS. Take 10 mL of the cells, add 80 mL D-PBS and 10 mL 0.5% trypan blue, mix and use this mixture to count the cells on a hemocytometer. Use one million cells for the CD19/LysoTracker assay. Mouse blood cells can also be cryopreserved for later analysis (see above).
4.3 B-LYMPHOCYTE STAINING 4.3.1 Materials and reagents for human cells PE-conjugated mouse anti-human CD19 antibody (clone LT19; Abcam) D-PBS (SigmaeAldrich) 15-mL centrifuge tubes 10% BSA FACS buffer (0.1% BSA, 0.02 M NaN3 in 1 PBS) 200 nM LysoTrackerÒ-green DND-26 (Invitrogen)
4.3.2 Protocol for human cells • • • • • • • • • •
Incubate one million mononuclear cells in a volume of 100 mL with 15 mL PE-conjugated mouse antihuman CD19 antibody in the dark on ice for 30 min. After 30 min, add 100 mL 10% BSA. Spin the cells for 5 min at 800 x g. Remove the supernatant. Resuspend the cells in 1 mL of 200 nM LysoTrackerÒ-green DND-26. Leave this in the dark at RT for 10 min. Spin the cells for 5 min at 800 x g. Remove the supernatant. Resuspend the cells in 500 mL FACS buffer and transfer the cells into FACS tubes. Put the cells on ice and analyze them on the FACS as soon as possible in a standardized way every time you run the assay (see below).
4.3.3 Materials and reagents for mouse cells Fc-block (0.5 mg/mL purified anti-mouse CD16/CD32; BD Pharmingen) CD19-PE (0.2 mg/mL PE anti-mouse CD19; BD Pharmingen) D-PBS (SigmaeAldrich) 15-mL centrifuge tubes 10% BSA FACS buffer (0.1% BSA, 0.02 M NaN3 in 1 PBS) 200 nM LysoTrackerÒ-green DND-26 (Invitrogen) FACS tubes
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4.3.4 Protocol for mouse cells • • • • • • • • • • •
Incubate one million cells in a volume of 50 mL with 2 mL Fc-block on ice for 5 min. Add 5 mL CD19-PE and incubate for 30 min in the dark on ice. Stop the incubation by adding 100 mL 10% BSA. Spin the cells for 5 min at 800 x g. Remove the supernatant. Resuspend the cells in 1 mL of 200 nM LysoTrackerÒ-green DND-26. Leave this in the dark at RT for 10 min. Spin the cells for 5 min at 800 x g. Remove the supernatant. Resuspend the cells in 500 mL FACS buffer and transfer the cells into FACS tubes. Put the cells on ice and analyze them on the FACS as soon as possible in a standardized way every time you run the assay (see below).
4.4 FLOW CYTOMETRY, CALIBRATION, ACQUISITION, AND ANALYSIS We use a BD Biosciences FACSCanto II for all our flow cytometric analysis, but other flow cytometer instruments can be used. Representative images showing the Fluorescence-activated cell sorting (FACS) analysis are shown in Figure 3. It is best however to use the same machine for all analysis if you need to compare the results over time as absolute values rather than as fold-changes (see discussion below).
4.4.1 Materials and reagents BD Biosciences FACSCanto II Cytometer Setup and Tracking beads (BD) Anti-mouse CompBeads (Anti-Mouse Ig, k/Negative Control (FBS) Compensation Particles Set, BD) 8-peak Rainbow calibration beads (SPHEROÔ Rainbow Calibration Particles, BD)
4.4.2 Protocol • • •
• •
Calibrate the cytometer using Cytometer Setup and Tracking beads (BD). Compensation is performed using cells stained with LysoTracker and anti-mouse CompBeads (BD) stained with PE antibody using FACSDiva software (BD). Samples are acquired with gating on singlet cells (FSC-H vs FSC-A) and CD19þ events. In total, 50,000 singlet events and 10,000 singlet gate CD19þ events are collected. The mean fluorescence of the CD19þ events is calculated using FACSDiva software (BD). The molecules of equivalent fluorescence (MEFL) is calculated using 8-peak Rainbow calibration beads (BD), using the fluorescein equivalent values provided by the manufacturer.
This shows singlet gating, Forward Scatter (FSC) versus Side Scatter (SSC) for human blood, the CD19 population identified with anti-CD19 antibody staining (red) and LysoTracker staining in green, and histogram of molecules of equivalent fluorescence (MEFL) of LysoTracker on B-cell population. (See color plate)
4. Methods
FIGURE 3 Representative images of flow cytometry setup windows.
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4.5 ISOLATION OF B-CELLS FOR BIOCHEMICAL ASSAYS OR MICROSCOPY When B-cells are needed for biochemical assays or for microscopy, they can be isolated from the mononuclear cell fraction isolated (as described above), using magnetic beads. We use magnetic cell separation (MACS) beads from Miltenyi Biotec for this purpose. We have used Dynabeads Mouse pan B (Life Technologies) in the past and found that these beads were fluorescent under a microscope, but are fine if biochemical analysis is to be performed, not microscopy.
4.5.1 Materials and reagents for human cells MACS buffer (D-PBS supplemented with 0.5% BSA and 2 mM EDTA) MACS Human CD19 MicroBeads (Miltenyi Biotec) 15-mL centrifuge tubes Tube roller bank (Miltenyi Biotec) MS column (Miltenyi Biotec) MACS separator (Miltenyi Biotec)
4.5.2 Protocol for human cells • • •
• • • • •
• •
Resuspend the mononuclear cells in 80 mL of MACS buffer per 10E7 total cells. Add 20 mL of MACS Human CD19 MicroBeads per 10E7 total cells, mix well, and incubate for 15 min on a tube roller bank at 4 C. Wash the cells by adding 10e20 x the labeling volume of buffer, centrifuge at 300 x g for 10 min, remove the supernatant completely, and resuspend the cell pellet in 500 mL MACS buffer. Place an MS column in the magnetic field of a MACS separator. Wash the column with 500 ml MACS buffer. Apply the cell suspension to the column. Rinse three times with 500 mL MACS buffer. Remove the column from the separator, place the column on a 15-mL tube, pipette 1 mL of MACS buffer onto the column, and flush out the positive cells using the plunger supplied with the column. Count the B-cells. Spin down the cells, remove the supernatant and wash them once with 1 mL D-PBS, remove the supernatant again and freeze the pellet at 80 C for biochemical analysis later or use the cells straightaway for staining for microscopy.
4.5.3 Materials and reagents for mouse cells MACS buffer (D-PBS supplemented with 0.5% BSA and 2 mM EDTA) MACS Mouse CD19 MicroBeads (Miltenyi Biotec) 15-mL centrifuge tubes Tube roller bank (Miltenyi Biotec) MS column (Miltenyi Biotec) MACS separator (Miltenyi Biotec)
4. Methods
4.5.4 Protocol for mouse cells • • •
• • • • •
•
Resuspend the mononuclear cells in 90 mL of MACS buffer per 10E7 total cells. Add 10 mL of MACS Mouse CD19 MicroBeads per 10E7 total cells, mix well, and incubate for 15 min on a tube roller bank at 4 C. Wash the cells by adding 10e20x the labeling volume of buffer, centrifuge at 300 x g for 10 min, remove the supernatant completely, and resuspend the cell pellet in 500 mL MACS buffer. Place an MS column in the magnetic field of a MACS separator. Wash the column with 500 mL MACS buffer. Apply the cell suspension to the column. Rinse three times with 500 mL MACS buffer. Remove the column from the separator, place the column on a 15-mL tube, pipette 1 mL of MACS buffer onto the column, and flush out the positive cells using the plunger supplied with the column. Spin down the cells, remove the supernatant and wash them once with 1 mL D-PBS, remove the supernatant again and freeze the pellet at 80 C for biochemical analysis later or use the cells straightaway for staining for microscopy.
4.6 INFLUENCE OF PATIENT BLOOD SHIPPING TIMES One of the practical considerations before implementing this method in your laboratory is to consider the effect of blood storage/shipping times on assay performance. Several practical scenarios can be envisaged. In a clinical setting, it may be possible to draw and analyze patient blood the same day in which case the situation is straightforward. However, even in this scenario it may not be possible to perform the assay on the same day depending on clinic visit times for the patient and so the question is: How should the sample be stored and does storage affect the results? This is even more of an issue in a research setting where samples will be shipped to the research laboratory for analysis from a clinic that may be in another country. This involves international shipments and considerable lag times between drawing the blood and their arrival in the research laboratory for analysis. The two key variables to consider are the blood sample storage/shipping temperature and its influence on assay performance and also how long a sample can be in transit and still be suitable for assay. We recently completed a longitudinal study spanning many years on Niemann-Pick type C patient blood samples and had to deal with these practical issues (te Vruchte et al. 2014). What we will do is share our experiences with you, so they can inform your choices for your own studies/own situation and show you some experimental data on how temperature and time affect the analysis. The simple take-home message is you must adopt one protocol and stick to it rigidly to allow accurate comparisons to be made between samples over time.
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4.7 INFLUENCE OF BLOOD STORAGE TEMPERATURE ON LYSOTRACKER STAINING Blood samples collected into EDTA vacutainer tubes can be stored at RT or can be refrigerated at 4 C. As our study (te Vruchte et al. 2014) required shipping of samples from Germany, the UK, and the USA, we defaulted to RT, as it was simpler for the clinics to deal with. The downside was that this was prone to changes in ambient temperature at airports depending on the time of year but generally speaking was in the 20e25 C range. Here we have run a small experiment to show you how temperature affects the assay and why you need to standardize this parameter. We drew blood from healthy volunteers and compared LysoTracker staining values (MEFL) in their B-cells when we stored the blood at RT or 4 C for up to 5 days. The samples were assayed daily as described above. The data are summarized in Figure 4. The samples maintained at 4 C were less affected by storage than RT samples, with the major reduction in MEFL signal occurring over the first 24 h. Interestingly, B-cells surviving out to 5 days albeit low in number gave very robust MEFL values comparable to B-cells assayed on day 1e2 (we since repeated this and made the same observation, not shown). As LysoTracker is a live cell stain, there is no need to worry about dead cells in this assay. The take-home message is as long as one temperature and one analysis time (days post-blood draw) are chosen, data over time and between individuals can be reliably compared. Fold change over time rather than
FIGURE 4 Effect of storing blood samples at 4 C or room temperature for 5 days. Samples were collected from five healthy volunteers and molecules of equivalent fluorescence (MEFL) of B-cells assayed by FACS on a daily basis for 5 days.
4. Methods
absolute values can also be used to compare data between centers. Flow cytometers also vary, so absolute comparisons will be difficult to make between instruments. Again, fold change relative to control samples can be used reliably.
4.8 INFLUENCE OF DELAYS IN ANALYSIS OF SAMPLES POST-LYSOTRACKER STAINING Another potential variable is the lag time between staining with LysoTracker and actually analyzing the samples on the flow cytometer. We evaluated this variable (Figure 5), and rapid analysis times gave the strongest signals. It is therefore important to standardize this carefully in any protocol you establish. Long delays give reduced sensitivity (low MEFL values), so analyzing within the first 30 min is recommended and should be kept constant from assay to assay.
4.8.1 Some economic considerations relating to B-cell staining The assay we describe is low cost (assuming you have a flow cytometer already in place) except for the LysoTracker and the anti-CD19 reagents. We initially followed manufacturer’s instructions for the anti-CD19 staining, but as you can see in Figure 6 we were able to titrate the antibody considerably and still achieve robust delineation of the CD19-positive population. So titrating the antibody you purchase can save
FIGURE 5 Effect of delaying analysis after LysoTracker staining. Analysis of samples within 30 min gave optimal signal intensity, which then declined up to 2 h post-staining. This illustrates the need to rapidly analyze samples after live cell staining with LysoTracker and the need to standardize the protocol, so this does not vary between experiments.
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FIGURE 6 Titration of anti-CD19 antibody. As can be seen, considerable titration of anti-CD19 was possible, well below the manufacturers recommended antibody concentration of 20 mL.
you money. The only requirement is that you use enough antibody to be able to identify the B-cells as a discreet population.
5. SUMMARY Relative lysosomal volume can be readily measured in circulating B-cells in mouse and human blood samples for studying LSDs. As long as you adopt a rigidly standardized protocol in relation to day of analysis post-blood draw, temperature the blood is stored/shipped at, and time from addition of LysoTracker to analysis on the cytometer, the assay is robust. We suggest that when comparing data between centers, where different cytometers/protocol times etc., are being used, a fold change from baseline or age-matched controls is the most pragmatic solution for comparing data rather than absolute values.
ACKNOWLEDGMENTS DtV and FMP were supported by Action Medical Research, Niemann-Pick UK and an unrestricted grant from Actelion. KLW is supported by a Stratified Medicine Grant (Gaucherite) from the MRC. FMP is a Royal Society Wolfson Research Merit Award holder. We thank the volunteers who donated blood for this study.
References
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