Application of Confocal Microscopy for

Microscopy: Science, Technology, Applications and Education A. Méndez-Vilas and J. Díaz (Eds.) ______________________________________________

Application of Confocal Microscopy for Quantification of Intracellular Mycobacteria in Macrophages P. Bettencourt, D. Pires, N. Carmo and E. Anes Mycobacteria-Host Interactions Unit. Centro de Patogénese Molecular - URIA, Faculdade de Farmácia and Instituto de Medicina Molecular, Universidade de Lisboa, Av. Prof. Gama Pinto, 1649-003 Lisboa, Portugal. This chapter describes the techniques used to prepare a uniform and consistent mycobacterial culture and for the infection of macrophages in vitro. Here, protocols are described for the achievement of a certain number of single cell bacilli per macrophage. Confocal microscopy in combination with the software ImageJ are highlighted, and these techniques will be correlated with quantification by FACS and confirmed by colony forming units (CFU) the classical method to validate the intracellular survival of Mycobacterium tuberculosis. Conventional CFU for quantification of intracellular slow growing mycobacteria is labour-intensive, with incubation requirements that can take up to several weeks. New alternatives and fast methods are required for a rapid assessment of the immune response as well to test new antibacterial drugs in highthroughput screens. Keywords Mycobacterium tuberculosis; Macrophages; Host-Pathogen Interactions; Confocal Microscopy;

1. Introduction Mycobacterium tuberculosis species complex (MTC), M. tuberculosis, M. bovis, M. africanum, M. canetti and M. microti, are the etiologic agents of tuberculosis, a major worldwide health problem [1, 2] . The success of these pathogens relies on their ability to inhibit host immune defences and persist in a potentially hostile environment, namely the macrophage phagosome by mechanisms not completely understood [3] . Furthermore, the increased incidence of multidrug-resistant strains brings back concerns in respect to the emergence of the disease and the urgent need to search for new effective antibacterial drugs [4] . There is a wide-range of growth characteristics among the genus Mycobacterium: from 3 hours of generation time in M. smegmatis, a non-pathogenic mycobacteria, to 18-24 hours in M. tuberculosis. This is particularly due to the extremely high content of complex lipids present in the cell wall, thus making it extremely difficult and challenging for efficient, uniform and reproducible infection experiments. Therefore mycobacteria, tends to form residual bacterial aggregates that are difficult to disperse as individual bacteria. The absence of bacterial clumps is important because these are rapidly trafficked to phago-lysosomes within macrophages, in contrast to phagosomes containing single M. tuberculosis bacilli [5] . Attaining precise quantitative and qualitative analysis of mycobacterial infections has been an ongoing challenge. Macrophage cell cultures in vitro are a suitable model to study mycobacteria-host interactions, however literature reports on the concept of multiplicity of infection (MOI) are unclear, confusing and misleading. The signalling for innate immune mechanisms is widely dependent, from the very early steps of infection, on the bacterial load per macrophage, and from the percentage of infected host cells. Profiting from mycobacteria expressing Green Fluorescent Protein (GFP), a vast array of recent technologies, based on fluorescence such as confocal microscopy or FACS are now being applied to better understand the host-pathogen mechanisms as well to test new drugs. Confocal microscopy, associated with freely available data analysis software, allows the precise evaluation of both quantitative and qualitative aspects of this model of infection. Although the qualitative evaluation of phenotypes depends, to a great extent in the specificity of the antibodies used, the quantitative analysis of intracellular pathogen survival in macrophages at discrete time points can be precisely determined using confocal microscopy.

2. The significance of multiplicity of infection on the context of mycobacterial infections In most experiments the multiplicity of infection (MOI) of 10 is used, with the concept of each host cell being infected with 10 bacteria. For this the vast majority of laboratories estimate the ratio mix per tissue culture (TC) well plate of one mammalian cell for 10 of bacteria. This does not take in account the source of the macrophage, the animal or bacterial species used or the virulence of different bacteria strains. Murine J774 macrophages internalize bacteria very efficiently in a short time compared with another, often utilized THP-1 human derived macrophage cell line [6, 7] . Virulent strains of M. bovis are internalized in higher amounts by monocytic blood bovine derived macrophages than M. bovis BCG [6] .

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The co-culture period of macrophages and bacteria that we describe here is called the pulse phase of the experiment. The time taken for bacteria infection as pulse /uptake is also an important factor to be considered. In our experiments, after 1 hour of pulse in J774 macrophages and M. smegmatis at an estimated MOI of 10, we obtain a homogenous infection of 1 to 3 bacteria per macrophage (Fig. 1a). This is much lower from the estimated rate and, in reality corresponds to a MOI of approximately 1. In contrast a pulse of 3 hours will lead to 5-10 bacilli per macrophage under our experimental conditions [7, 8] . However, if one is expecting to follow an early signalling event, such as NF-κB activation or a MAP kinase phosphorylation, by using this bacterial load, 3 hours post-bacteria uptake may be ain inappropriate time to study these early events [7, 9, 10] . Indeed, to reach 5 to 10 of M.smegmatis load per macrophage from the J774 lineage, at early time post- uptake we needed to used an estimated MOI of 100 [7] . Therefore it is mandatory for each set of experiments to assess the real number of bacilli obtained per macrophage for each given pulse time, through confocal microscopy. Depending on the real bacterial load per cell, distinct proinflammatory events and cell signalling cascades will be displayed and, at later time points accurate numbers of intracellular bacterial survival will be obtained. Another important issue on the context of mycobacterial infections is the high content in complex glycolipids of the cell wall, making it extremely difficult and challenging to produce an efficient, uniform and, reproducible singlebacteria suspension. In fact bacterial clumps are rapidly trafficked to phago-lysosomes within macrophages, in contrast to phagosomes containing individual M.tuberculosis bacilli [5] . 2.1 Single Bacterial Suspension preparation In order to avoid the formation of residual bacterial aggregates that are difficult to disperse as individual bacteria (Fig. 1b), our laboratory is using the following protocol: Usually, mycobacteria grows in medium containing Middlebrook’s 7H9 broth Medium (Difco), Nutrient broth (Difco) (4.7 g and 5 g per liter, respectively), supplemented with 0.5% glucose for the fast growing M. smegmatis, or Middlebrook’s 7H9 broth Medium supplemented with OADC (oleic acid, albumin, dextrose and catalase) 10% from Difco, for slow growers such as M. tuberculosis or BCG. The cultures should be grown on a shaker at 220 r.p.m and at 37°C. Tween 80 at 0.05% could be added but some authors claim that some lipidic virulence components will be lost. Bacterial cultures in exponential growth phase are pelleted at 200 g, for 10 min. washed twice in PBS pH 7.4 and ressuspended in DMEM or other appropriate cell culture medium. The suspension is kept on the bench for 5 minutes, to allow the decantation of the large clumps of bacteria. Clumps of bacteria are removed by ultrasonic treatment of bacteria suspensions in an ultrasonic water bath for 15 min. The suspension is then passed through a 5 µm pore filter membrane. Single cell suspension is confirmed by phase contrast microscopy or in a fluorescence microscope (if bacteria are fluorescent). In the absence of clumps, the OD of the suspension at λ = 600 nm should then be adjusted to 0.1. We generally assume that 0.1 OD600 corresponds to 1X107 bacteria per ml. However, we strongly recommend that this suspension should be plated and counted by CFU, to more accurately determine the total number of bacteria, as this can vary immensely as a function of the growth medium composition of mycobacteria. Pitfalls and troubleshooting: in the case of bacteria remaining in clumps, use a 5 ml syringe needle (27 or 28 gauge) to disrupt the clumps. Collecting the complete volume of a suspension and pressing it against the tube wall, at least 30 times should be sufficient to get individualized bacilli. 2.2 Infection procedures Upon dilution of the suspension to a desired number of bacteria, the infection of host cells can proceed. Adherent host cells should be previously counted and transferred to a 6-, 24- or 96- TC well plate and, at the time of infection, cells should have achieved approximately 80% confluence. Large empty spaces on the well should be avoided because noninternalized bacteria by phagocytes may attach on the plastic surface and remain there even though a number of washing steps are applied. Conversely, if the number of host cells start to exceed the capacity of the well, the cells will exhaust the medium nutrients very rapidly and will become round-shaped and activated due to the close contact with neighbor cells. Finally, small round-shaped cells will loose the capacity to phagocyte bacteria compared to enlarged cells with free space around them. Overall, the area of empty spaces in an infection of adherent cells should be tightly controlled. Using an estimated MOI of 10 (meaning then fold higher amounts of bacteria than macrophages in a tissue culture plate mix) we can obtain a fairly good and reproducible infection rate of total 70-80% infected macrophages having after a 1 hour pulse 1 to 3 bacteria per host cell. The co-culture of macrophages and bacteria is called the pulse phase of the experiment. Removing the bacteria and washing the macrophages brings the experiment to the second part, the chase of the experiment. In pulse-chase experiments, when infecting adherent host cells, the MOI does not take in account the volume of the suspension to be used. To avoid this uncertainty, many authors centrifuge the bacterial suspensions immediately after transferring it to the well containing the adherent host cells. This is performed at low speed in order to increase the number of bacteria in direct contact with the host cells. We do not perform this centrifugation step due to the fact that in

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order to spin down all the bacteria in any suspension it is necessary to spin at 200 x g. At this speed the host cells would be dramatically damaged. On the other hand, using lower spin speeds of around 120 g, which are commonly used, would only spin down the remaining large aggregates of bacteria (if present), leaving most of the individual bacteria floating in the suspension. Therefore, centrifugation is unnecessary. Furthermore, using centrifugation to enhance the contact between bacteria and host cells introduces a new element to the experiment: the centrifugal force. In some internalization assays this artificial force would be deleterious for the host cells that might be particularly sensitive to external forces. Therefore the use of centrifugation to enhance the contact between bacteria and host cells should be strongly avoided. To minimize the interference from the extracellular fluid, we use a minimum volume of infection medium (the medium used to grow host cells, without antibiotics). This volume has to obey a few rules including covering every host cell with medium, during the infection period. For the chase part of the experiment, the volume is adjusted depending on the size of the TC well plate. 2.3 Immunofluorescence After the infection experiment, cells and bacteria can literally be “frozen” for visualization. Cell fixation will immobilize all the organelles and sub-cellular structures at the time of the treatment. Although fixation with agents such as paraformaldehyde can be a harsh process, specific intracellular components can be visualized after staining or labelling of desired host or parasite molecules located at distinct subcellular structures. Although the qualitative evaluation of phenotypes depends, to a great extent on the specificity of the antibodies used, the quantitative analysis of intracellular pathogen survival within macrophages can be precisely determined using confocal microscopy, at discrete time points. The protocol we use for immunofluorescence is detailed bellow: Transfer a suspension of the desired number of adherent host cells on top of glass coverslips placed inside the holes of a 24-well plate. This should be done at least one day before the experiment to allow the cells to attach to the coverslip. At the desired time point to observe the cells, the medium should be removed and the cells washed twice with PBS, at room temperature. Fix the cells at room temperature for 30 mins with PBS containing 4% w/v PFA and 4% w/v sucrose. After fixation, all subsequent steps can be performed on the bench. All reagents should be used at room temperature. Wash the cells twice with PBS. Incubate with NH4Cl 50 mM in PBS for 15 minutes. Wash the cells once with PBS. To permeabilize the fixed cells, 0.1% w/v triton in PBS for 5 minutes. Wash the cells twice with PBS. Incubate the cells with 1% w/v bovine serum albumin (BSA) in PBS for 15 minutes. Incubate the cells with the desired antibodies, at specified temperature, diluted in the 1% BSA solution for 30 minutes. Wash the cells twice with PBS and mount the cover-slip face down on to a cover slide containing mounting medium. As we use oil immersion lenses, we recommend the usage of glycerol based mounting media containing antifade agents, such as Dako or Vectashield.

3. Quantitative Data Analysis Using Confocal Microscopy Confocal microscopy is one of the highest resolution light microscopy methods available. Using 63X objectives (Zeiss) with very high optical aperture and oil immersion, one can reach a maximum resolution of approximately 0,2 µm in XY axis and 0.7 µm in XZ axis. In our particular setting, the smallest and most important object to observe are the individual GFP expressing bacteria, which are around 2 µm long and 0.2 µm of diameter. Therefore, it is possible to detect individual bacteria with a reasonable resolution, using confocal microscopy. Studying sub-cellular organelles may be a greater challenge as fixation conditions are harsh and may destroy delicate intracellular structures thus giving a false impression of the morphology of the structures housing the bacteria. However, to quantify the number of individual bacteria inside one host cell, this technique is extremely powerful. Using particular experimental settings, mycobacteria can be highly internalized by host cells. And when this is the case, the laborious and time consuming process of manual counting of hundreds of bacteria per host cell can be circumvented by the use of imaging software ImageJ, a free application for microscopy data analysis that is currently being used by most people in the field

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(http://rsbweb.nih.gov/ij/). ImageJ also allows one access to thousands of specific plugins, developed by labs all over the world, to address specific questions and applications. Therefore, when doing image analysis, it will be very easy to find someone on the web that will have developed a particular application to fulfill your needs. If this is not the case, one can create its own macros to analyze the most demanding microscopy images. Finally, ImageJ will read and process any image in any format, independently of the software used for its creation. In our lab we developed a macro to quantify every single bacteria on host cells even if they are aggregated in clumps. Simultaneously this macro can count all the host cells. Using this macro, which is freely available on request (http://www.ff.ul.pt/paginas/eanes/Site/Research.html ), one can quantify the infection rate automatically in a fraction of time it would take to do this via manual counting. The quantification can be made in a single focal plan, although this will give a biased result, as bacteria can be dispersed throughout the host cell. To circumvent this, one can acquire multiple images in different focal planes to create a Z-stack of images (data) through the Z axis. A correctly acquired Z-stack will have the complete number of bacteria inside the host cell. To quantify it automatically, a projection of all Z-stacks into one image has to be made. This image can be analyzed by our macro to give a very accurate count of the number of bacteria per cell.

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Fig. 1 Confocal microscopy image of J774 macrophages infected with M. smegmatis–GFP after 1h of pulse. Red: actin labeled using Rhodamine-Phalloidine staining; Green – Green Fluorescent Protein expressing bacteria a) Macrophages containing unicellular bacteria b) Macrophages containing clumps of bacteria.

4. Assessment of the infection: complementary assays for mycobacteria within macrophages 4.1 Micro colony-forming units assay The colony-forming units (CFUs) assay is one of the oldest and mostly used methods in microbiology to quantify bacteria in a sample. In our laboratory, the main use for this technique is to quantify the bacterial survival within host cells such as macrophages and dendritic cells. The assumption supporting this technique is that each bacteria plated on broth medium will form an individual colony. Thus, if we plate a sample of the suspension we want to analyse and count the number of colonies produced we can estimate the actual number of bacteria present in the original suspension. It is worthwhile to mention that several factors can lead to inaccuracies using this method. Firstly this technique will not measure all the viable bacteria inoculated but only those able to grow in the conditions that we cultivate them. For example latent M. tuberculosis though viable will not proliferate under these conditions. This is the reason why the results obtained with this technique for the quantification of a suspension of bacteria are written as “CFU/ml” and not “bacteria/ml”. Still, our advances in bacteriology have enabled us to maximize the number of bacteria able to produce colonies in a solid agar medium. For the case of mycobacteria, standard Middlebrook 7H10 medium is widely used to cultivate and access the CFUs. In the specific case of fastidious slow growing mycobacteria such as M. tuberculosis or M. bovis, the 7H10 medium is supplemented with OADC or ADC as previously described. Another important factor when considering this technique is that if an accurate number of CFUs in a suspension is required there should be emphasis in making that bacterial suspension as much evenly distributed as possible. This will

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ensure that the plated sample is representative of the original suspension. Also, if two bacterial cells are plated next to each other they will grow and merge resulting in the formation of one single colony and thus hindering the true number of bacteria actually plated. Mycobacteria have a tendency to form aggregates due to the lipidic content of their cell wall. In this protocol we highlight the use of detergents to prevent bacterial aggregation while the bacteria grow in broth medium such as Middlebrook 7H9. While in most quality control tests the number of bacteria present in the analysed sample is expected to be low, that is usually not the case for infection assays. In most of our experiments with mycobacteria and macrophages in 6-, 24- or 96-well plates, the MOI used is 10 bacteria per macrophages. This leads to a final number of internalized bacteria around 104-105 bacteria in each well plate. To quantify these samples there is need to perform serial 1/10 dilutions thus increasing the error in the measure and one can easily understand how troublesome this technique is. Performing CFU assays for testing numerous amounts of conditions is both expensive and time consuming. Slow growing bacteria such as M. tuberculosis take about 20-24 h to divide. On a plate, a single individualized bacterium will take 3-4 weeks to generate a visible colony. This means that any assay performed today will only generate results after a month of incubation. Moreover, the standard deviation associated with this technique (20% of the mean when performed by experienced personnel) diminishes the utility of this method for identifying subtle changes in bacterial survival. Nevertheless, in spite of the disadvantages of using this technique it remains the most reliable and widely accepted. When developing or implementing other quantitative methods, CFU counting is the standard we have to dethrone and so far, that remains to be done. The micro colony-forming units assay described below is based on the original CFU method to count host cellinternalized viable bacteria with minor modifications. Colonies are counted from small drops of the suspension instead of using the whole agar plate. This is a small step to maximize the resources since a single agar plate may hold several drops for counting CFU. Another advantage is that we can count the colonies in these small areas by using a simple microscope. This reduces the time one would normally need to have results visible by eye. The CFU protocol is thus: Remove cell culture medium and wash the cells 1x with PBS. Add Igepal 1% (or water) (100 µl, 500 µl or 1000 µl for 96-, 24-, or 6-well plates, respectively) in order to lyse the host cells but not the bacteria. Incubate 10-15minutes at 37ºC. Mix the suspension with the pipette several times while scratching the bottom of the plate. Take 20 µl of the suspension and make serial 1/10 dilutions in water on a 96-well plate to a final volume of 200 µl (20 µl+180 µl). Usually we perform 4 dilutions (10-1, 10-2, 10-3, 10-4), but if using a low MOI, one could consider plating directly from the lysate and performing only 3 dilutions (10-0, 10-1, 10-2, 10-3). To ensure more reliable results you may dilute each sample twice and separately to have technical duplicates. A

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For a solid culture plate for mycobacteria (7H10 medium prepared and supplemented according to the manufacturer), a drawing scheme of the dilutions is shown in Fig. 2. It is important that the medium is thick enough (20-25 ml) if testing slow growth mycobacteria since the plates will be in the incubator for 2-3 weeks. Whilst the plates must endure for a long time in the incubator you should let them dry before using them. This will prevent surface humidity from spreading the suspension drops. Pipette a 10 µl drop of the diluted suspensions on each dot on the plates’ scheme. If you used 1% Igepal solution to lyse the cells then you may consider plating 5 µl instead of 10 µl since the detergent tends to spread the drops. Let the drops dry and incubate the plates at 37ºC. Make sure the incubator is fully supplied with water to prevent the plates from drying completely. After 24 h to 48 h for fast growth mycobacteria or 2-3 weeks for slow growth mycobacteria colonies will be visible trough a microscope or a magnifying glass.

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From those colonies we can estimate the number of bacteria in the original lysate. Results can be calculated and represented as CFU/ml or CFU/sample(well) (Equation 1). Equation 1.

CFU / sample = number _ of _ colonies ×

1 dilution _ factor

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Analysing the results from several experiments is troublesome and time consuming and some of the methods implemented to ease this process include automated colony counters. Several types of apparatus for performing this are available in the market and these automatically scan the agar plates for unstained, stained or even fluorescent bacterial colonies. Other “homemade” apparatus are also possible such as using digital cameras or scanners. Here an image of the plate can initially be captured and then processed further using free software packages such as ImageJ this already possess the needed macros that would allow one to automatically count colonies on a picture. 4.2 Quantification of mycobacteria internalization by FACS. Fluorescence activated cell sorter (FACS) allows the observation of population specific events. When applied to bacteria internalization assays by host cells, combining mycobacteria labeled with fluorescent reporters, it enables one to quantify on a cell-to-cell basis the percentage of cells containing intracellular mycobacteria. Although FC is a powerful technique for quantification of fluorescent intracellular mycobacteria in macrophages, it cannot distinguish between internalized and surface bound bacteria. To discriminate this, fluorescent from surface bound bacilli should be quenched. Alternatively the use of fluorescent labeled antimycobacterial antibodies that reaches external but not internalized bacteria can be used. In our laboratory we are using Green Fluorescence Protein (GFP) expressing Mycobacterium to perform pulse-chase internalization experiments by macrophages. To utilize FACS in this context we use the following protocol: Take 10 ml of an exponentially growing mycobacterial culture and spin 200 x g for 10 minutes. Wash with 10 ml of warmed PBS and spin at 200 x g for 10 minutes. Resuspend in 5 ml of the appropriate cell culture medium. Measure Optical Density at 600 nm and calculate the desired mycobacterial concentration for infection (OD600=0,1=107 mycobacteria/ml) to achieve the desired MOI in 50 µl volume. Remove the medium from the cells, add the mycobacteria and incubate for a 1hour pulse. Remove the infection media and wash the cells with 200 µl of warmed PBS three times. Trypsinize the cells by adding 50 µl of trypsin 0,25 % , incubate for 5 minutes and ressuspend in 150 µl of cell medium. Transfer the cells to a 96 round bottom well plate, spin at 120 g for 5 minutes. Carefully recover the supernatant, and ressuspend the cells in 200 µl of PBS and spin at 120 g for 5 min. Repeat the last procedure and add 200 µl of PBS with 10% of FCS (fetus calf serum). Pass cells through the FACS and analyse the GFP positive cells.

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Fig. 3 Confocal microscopy image of J774 macrophages pulsed with carboxyfluorescein coated beads and quenched with trypan blue. External beads are quenched within 8 min (white circles). Fixation of cells with PFA will reverse the quenching.

Using the FC analysis program FlowJo, the analysis starts by gating in the Forward Scater (FSC) and Side Scater (SSC) in the sample (Fig.4a and 4c). The gated population can be used to make a SSC /FL1 and distinguish between THP-1 cells that internalized GFP expressing bacteria (GFP positive) (Fig. 4d) and bacteria-free cells (Fig. 4b). Transforming the FL1 data in histograms one can compare the distribution of fluorescent intensity in the GFP positive population. 10 4

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Fig. 4 FACS analysis of THP-1 macrophages using FlowJo software. (A and C) Macrophage sample acquired in the Forward Scater (FSC) and Side Scater (SSC). (B) The cell population can be subsequently gated and a precise quantification of the macrophages that do not contain GFP-bacteria and (D) those who contain GFP-bacteria, at a MOI of 20. E) Different amounts of bacteria are distinguishable as a function of the fluorescence intensity. GFP-positive cells show higher intensity as the MOI increases from 5 (blue), to10 (green) and to 20 (yellow).

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Acknowledgements The support from the national agency Fundação para a Ciência e Tecnologia (projects PTDC/ BIABCM/102123/2008; PTDC/SAU-MII/098024/2008, and PIC/82859/2007) is gratefully acknowledged. We are also grateful to Maximiliano Gutierrez, Helmholtz Centre, Braunschweig, Germany and to Arwyn Tomos Jones, Welsh School of Pharmacy, Cardiff University for critical reading of this manuscript. PB, DP and NC are FCT PhD fellows. We are grateful to Nuno Moreno from the Instituto Gulbenkian de Ciência for technical support in developing the ImageJ macros.

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