Fluorescence Microscopy Methods for Determining the Viability of Bacteria in Association with Mammalian Cells

Summary

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. This article outlines protocols for the use of fluorescent dyes that reveal the viability of individual bacteria inside and associated with host cells.

Abstract

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. Current protocols to address these questions include colony count assays, gentamicin protection assays, and electron microscopy. Colony count and gentamicin protection assays only assess the viability of the entire bacterial population and are unable to determine individual bacterial viability. Electron microscopy can be used to determine the viability of individual bacteria and provide information regarding their localization in host cells. However, bacteria often display a range of electron densities, making assessment of viability difficult. This article outlines protocols for the use of fluorescent dyes that reveal the viability of individual bacteria inside and associated with host cells. These assays were developed originally to assess survival of Neisseria gonorrhoeae in primary human neutrophils, but should be applicable to any bacterium-host cell interaction. These protocols combine membrane-permeable fluorescent dyes (SYTO9 and 4',6-diamidino-2-phenylindole [DAPI]), which stain all bacteria, with membrane-impermeable fluorescent dyes (propidium iodide and SYTOX Green), which are only accessible to nonviable bacteria. Prior to eukaryotic cell permeabilization, an antibody or fluorescent reagent is added to identify extracellular bacteria. Thus these assays discriminate the viability of bacteria adherent to and inside eukaryotic cells. A protocol is also provided for using the viability dyes in combination with fluorescent antibodies to eukaryotic cell markers, in order to determine the subcellular localization of individual bacteria. The bacterial viability dyes discussed in this article are a sensitive complement and/or alternative to traditional microbiology techniques to evaluate the viability of individual bacteria and provide information regarding where bacteria survive in host cells.

Introduction

There is a dynamic interaction and co-evolution between bacteria and the hosts in which they reside. Bacteria have evolved adherence organelles, secretion systems, and/or the ability to produce toxins that enable their productive infection of host phagocytic and non-phagocytic cells. The bacteria must also contend with recognition and antimicrobial activities of the host immune system. The host immune system is comprised of innate and adaptive components including physical and chemical barriers, immune cells, the complement system, and other components of humoral immunity. While many bacteria are susceptible to killing and clearance by the multilayered host immune response, some pathogenic and opportunistic bacteria have evolved mechanisms to infect a variety of host cells and subvert clearance by the host immune response ¹. Neisseria gonorrhoeae is one example of a bacterial pathogen that is highly adapted to persist in its human host. N. gonorrhoeae readily colonizes the luminal surfaces of mucosal epithelial cells of the urogenital tract, pharynx, conjunctiva, and rectum. Colonization triggers the abundant recruitment of neutrophils at mucosal sites. Neutrophils are professional phagocytes that possess a variety of antimicrobial processes to kill microorganisms; however, N. gonorrhoeae is capable of surviving in the presence of neutrophils ^2-5. Understanding how bacterial pathogens such as N. gonorrhoeae subvert, suppress, and hijack the immune response to ultimately survive in normally hostile host environments is crucial to the development of new therapies for combating infectious diseases.

Experimental protocols often used to investigate bacterial survival in host cells include colony count assays, gentamicin protection assays, and electron microscopy. In colony count assays, a population of infected cells is lysed (for instance, with a detergent to which the bacteria are resistant) to liberate the bacteria. The lysates are diluted and plated on agar-based media, and colony-forming units in the lysates are enumerated for each time point and/or experimental condition. This approach reports the viability of the entire bacterial population but is not capable of differentiating between intracellular and extracellular survival. A variation on the colony count assay, the gentamicin protection assay, specifically measures intracellular bacterial survival, based on the inability of the antibiotic gentamicin to cross the eukaryotic plasma membrane ⁶. However, this assay is dependent on the bacteria being susceptible to killing by gentamicin (or another antibiotic that is similarly eukaryotic membrane-impermeant) and the inability of the antibiotic to have access to internal bacteria. Therefore, a gentamicin protection assay may not be effective for examination of all bacterial species or when examining bacterial survival in highly pinocytic cells such as neutrophils. Neither of these approaches reveals the subcellular localization or other behavior of individual bacteria (e.g. if the bacteria form aggregates or microcolonies that behave differently from individual bacterial cells). Another frequently used approach to examine the viability of individual external and internal bacteria is thin-section transmission electron microscopy (TEM). This approach is advantageous in that it can provide information regarding the location of the bacteria in host cells (e.g. phagosome, cytoplasm, autophagosome), which can be further investigated by immunoelectron microscopy with gold-coupled antibodies against subcellular markers. However, electron microscopy is not especially sensitive at assessing bacterial viability. When embedded sections are stained with uranyl acetate, lead citrate, or other electron-dense reagents and imaged by electron microscopy, electron-dense bacteria are considered viable and electron-lucent nonviable ^7,8. However, this assumption overestimates bacterial viability, since only those dead bacteria with severely disrupted membranes and devoid of cytoplasm appear electron-lucent. In addition, some bacterial species may display a range of electron densities depending on their stage of growth, making it difficult to determine viability.

As an alternative or in addition to these widely used methods, here we provide protocols and rationale for the use of fluorescent dyes that indicate bacterial viability to assess the survival of bacteria attached to and internalized by host cells. To identify extracellular bacteria, infected cells are first exposed to a fluorescent reagent, such as a lectin or bacteria-specific antibody. The infected cells are then permeabilized and exposed to DNA-specific dyes that are differentially accessible to bacteria with intact vs. degraded membranes, as a surrogate for bacterial viability. In the first protocol, the membrane permeable dye SYTO9 identifies the total bacterial population, while propidium iodide is only accessible to those bacteria that have compromised membranes and are thus considered nonviable. Propidium iodide and SYTO9 have been used to evaluate bacterial viability in biofilms, discriminate pathogenic from nonpathogenic bacteria, and enumerate viable water-borne bacteria ^9-12. In the second protocol, 4',6'-diamidino-2-phenylindole (DAPI) identifies total bacteria, while SYTOX Green is only accessible to the nonviable population. These viability dye pairs can be combined with immunofluorescence to determine each bacterium's location in relation to a protein of interest, for instance to define bacterial subcellular localization. The use of these assays provides key insight into the interactions that result in bacterial killing or survival during infection of host cells. The protocols outlined in this article were used to assess the viability of N. gonorrhoeae that is attached to and inside primary human neutrophils, including in different populations of neutrophil phagosomes ^5,13,14. However, these protocols can be applied to assess viability of gram-positive and gram-negative bacteria in professional phagocytes, non-professional phagocytes, and protozoa ^15-24.

Protocol

1. Assessing Bacterial Viability with Propidium Iodide and SYTO9

1. Infect cells that are adherent to 12 mm diameter circular glass coverslips in 24-well plates with bacteria of interest. Do NOT fix the cells with aldehydes or organic solvents.
2. Rinse cells once gently in 0.1 M 3-(N-morpholino)propanesulfonic acid (MOPS), pH 7.2, containing 1 mM MgCl₂ (MOPS/MgCl₂).
3. Incubate cells for 10 min in the dark at room temperature with Alexa Fluor 647-coupled antibody or lectin that binds to the bacterial species of interest, in MOPS/MgCl₂, to detect external bacteria.

NOTE: Conduct controls with live and dead bacteria, in the absence of host cells, to show that the lectin or antibody of interest binds all bacteria regardless of viability (Figure 1).

4. Rinse cells two times with MOPS/MgCl₂.
5. Aspirate media from cells and add 0.5 ml Live/Dead Staining Solution. Live/Dead Staining Solution is 5 μM SYTO9, 30 μM propidium iodide, and 0.1% saponin (final concentrations) in MOPS/MgCl_2.
6. Incubate cells for 15 min at room temperature in the dark.
7. Rinse cells two times in MOPS/MgCl_2.
8. Invert coverslips face down onto glass slides and seal with clear nail polish. Do not use mounting media.
9. Acquire images within 30 min, using a fluorescence microscope with filter sets compatible with green, red, and far-red image acquisition.

NOTE: Both conventional fluorescence microscopy and confocal laser scanning microscopy can be used. After 30 min the fluorescent dyes begin to leak from the bacteria and the data acquired are no longer accurate. Images shown in this article were acquired on a Nikon Eclipse E800 upright fluorescence microscope with Hamamatsu Orca-ER digital camera using Openlab software. Fluorescence of Alexa Fluor 647 was detected using a filter with excitation wavelength of 590 - 650 nm and an emission filter of 663 - 735 nm, and is false-colored blue. Fluorescence from propidium iodide was detected using a filter with excitation wavelength of 540 - 580 nm and an emission filter of 600 - 660 nm. Fluorescence from SYTO9 was detected using a filter with excitation wavelength of 465 - 495 nm and an emission filter of 515 - 555 nm.

10. This protocol will result in images where external nonviable bacteria appear blue + red, internal nonviable bacteria appear red only, external viable bacteria appear blue + green, and internal viable bacteria appear green only. Count by eye the numbers of bacteria that are external nonviable, internal nonviable, external viable, and internal viable.
11. Calculate the percent of external viable bacteria by dividing the number of external viable bacteria by the total number of external bacteria (viable plus nonviable). Calculate the percent of internal viable bacteria by dividing the number of internal viable bacteria by the total number of internal bacteria (viable plus nonviable) (Figure 4).
12. There are two essential controls to perform with this protocol. First, validate that all nonviable bacteria are propidium iodide-positive and all live bacteria are SYTO9-positive. A mid-logarithmic culture of bacteria (in the absence of any host cells) should be >95% SYTO9-positive. Shown in Figure 2 is a mid-logarithmic culture of N. gonorrhoeae (at 10⁸ colony forming units per ml) incubated with Live/Dead Staining Solution. Second, validate that both propidium iodide and SYTO9 can enter permeabilized, infected host cells.
    1. To generate a population of dead bacteria, collect 2 x 10⁸ bacterial colony forming units in a 1.5 ml microfuge tube, add 70% isopropanol and sit for 10 min. Pellet the bacteria in a microfuge and wash the bacteria twice in PBS to remove residual isopropanol. Then follow protocol steps 1.5 - 1.7. Add 5 μl of bacterial suspension to a glass slide and overlay with a coverslip. Image samples as in protocol step 1.9. Under these conditions, 100% of the population should be propidium iodide-positive (Figure 2). In some bacterial species, propidium iodide may not completely overwhelm SYTO9 staining, and nonviable bacteria may appear yellow or orange.
    2. For phagocytic cells like neutrophils, expose the cells to isopropanol-killed bacteria and ensure that 100% of the intracellular bacteria are propidium iodide-positive (Figure 3). For nonphagocytic cells that may not internalize dead bacteria, it may be sufficient to treat infected cells with sodium azide or other cell-permeant antimicrobial agents prior to adding Live/Dead Staining Solution.

2. Assessing Bacterial Viability with SYTOX Green and DAPI

1. Label bacteria of interest with 10 μg/ml DAPI in Morse's Defined Medium ²⁵ for 20 min at room temperature in the dark.
2. Infect cells that are adherent to 12 mm diameter circular glass coverslips in 24-well plates with DAPI-labeled bacteria. Do NOT fix the cells with aldehydes or organic solvents.
3. Rinse cells once with MOPS/MgCl_2.
4. Incubate cells for 10 min at room temperature in the dark with Alexa Fluor 647-coupled antibody or lectin that binds to the bacterial species of interest, in MOPS/MgCl₂, to detect external bacteria. See note in protocol step 1.3 for suggested controls.
5. Aspirate media from cells and add 0.5 ml 0.4 μM SYTOX Green in MOPS/MgCl₂.
6. Incubate cells for 5 min at room temperature in the dark.
7. Rinse cells two times in MOPS/MgCl₂.
8. Wash cells one time in MOPS/MgCl₂ for 5 min.
9. Acquire images within 30 min with a fluorescence microscope. See protocol step 1.9 for description of microscope, digital camera, and acquisition software.

NOTE: Fluorescence from Alexa Fluor 647 was detected using a filter with excitation wavelength of 590 - 650 nm and an emission filter of 663 - 735 nm, and is false-colored red. Fluorescence from SYTOX Green was detected using a filter with excitation wavelength of 465 - 495 nm and an emission filter of 515 - 555 nm. Fluorescence from DAPI was detected using a filter with excitation wavelength of 355 - 375 nm and barrier filter of 400 nm.

10. This protocol will result in images where external nonviable bacteria appear red + green, internal nonviable bacteria appear green + blue, external viable bacteria appear red + blue, and internal viable bacteria appear blue only (Figure 4). Quantify the percent of external and internal bacteria as described in protocol step 1.11.
11. The controls described in protocol step 1.12 should be performed with the DAPI/SYTOX Green dye combination (Figures 2 and 3).

3. Assessing Bacterial Viability Alongside Subcellular Localization

1. Label bacteria of interest with 10 μg/ml DAPI in Morse's Defined Medium for 20 min at room temperature in the dark.
2. Infect cells that are adherent to 12 mm diameter circular glass coverslips in 24-well plates with DAPI-labeled bacteria. Do NOT fix the cells with aldehydes or organic solvents.
3. Rinse cells once with MOPS/MgCl_2.
4. Incubate 10 min at room temperature in the dark with Alexa Fluor 647-coupled antibody or lectin that binds to the bacterial species of interest, in MOPS/MgCl₂, to detect external bacteria. See note in protocol step 1.3 for suggested controls.
5. Aspirate media from cells and rinse cells two times in MOPS/ MgCl_2.
6. Incubate cells with Alexa Fluor 555-coupled antibody against subcellular marker of interest for 20 min, in MOPS/MgCl₂ containing 0.2% saponin.
7. Rinse cells two times in MOPS/MgCl₂.
8. Wash cells one time in MOPS/MgCl₂ for 5 min.
9. Aspirate media from cells and add 0.4 μM SYTOX Green in MOPS/MgCl₂.
10. Incubate cells 5 min at room temperature in the dark.
11. Rinse cells two times in MOPS/MgCl₂.
12. Wash cells one time in MOPS/ MgCl₂ for 5 min.
13. Acquire images of slides within 30 min on fluorescence microscope. See protocol step 1.9 for description of microscope, digital camera, and acquisition software.

NOTE: Fluorescence from Alexa Fluor 647 was detected using a filter with excitation wavelength of 590 - 650 nm and emission filter of 663 - 735 nm, and is false-colored purple. Fluorescence from Alexa Fluor 555 was detected using a filter with excitation wavelength of 540 - 580 nm and an emission filter of 600 - 660 nm. Fluorescence from SYTOX Green was detected using a filter with excitation wavelength of 465 - 495 nm and an emission filter of 515 - 555 nm. Fluorescence from DAPI was detected using a filter with excitation wavelength of 355 - 375 nm and barrier filter of 400 nm.

14. This protocol will result in images where external nonviable bacteria appear purple + green, internal nonviable bacteria appear green + blue, external viable bacteria appear purple + blue, internal viable bacteria appear blue only, and subcellular protein appears red (Figure 4). Quantify the perecent of external and internal viable bacteria as described in protocol steps 1.11.
15. The controls described in protocol step 1.12 should be performed with the DAPI/SYTOX Green dye combination (Figures 2 and 3).
16. In addition to counting viable and nonviable bacteria, classify each bacteria as either positive or negative for colocalization with the subcellular marker of interest.
17. Calculate the percent of viable bacteria colocalized with the subcellular marker by dividing the number of colocalized viable bacteria by the total number of viable bacteria. Calculate the percent of nonviable bacteria colocalized with the subcellular marker by dividing the number of nonviable colocalized bacteria by the total number of nonviable bacteria.

Results

The protocols outlined were used to examine survival of N. gonorrhoeae after exposure to primary human neutrophils ^5,26. Neutrophils were infected with N. gonorrhoeae and processed with protocol 1, using the green-fluorescent viability dye SYTO9 and the red-fluorescent propidium iodide (Figure 4A). The dyes were added in the presence of saponin, which sequesters cholesterol to preferentially permeabilize host cell plasma membranes, not N. gonorrhoeae membranes. Other...

Discussion

Presented here are two protocols that use DNA binding and viability dyes in conjunction with a fluorescent lectin to identify live and dead bacteria attached to and inside human cells. Since both protocols effectively discriminate live from dead bacteria, the choice of which protocol to use depends upon the goal of the experiment. The first protocol uses propidium iodide to detect nonviable bacteria and SYTO9 to detect intact bacteria. Shown in Figure 4A is a representative image of protocol 1 applied to...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
21 G 3/4 butterfly needles for blood collection	Becton Dickinson	367251
Blood collection tubes with Sodium Heparin 10 ml	Becton Dickinson	366480
Sterile water for irrigation	Baxter	07-09-64-070
Dextran 500	Sigma	31392
Sodium Chloride	Fisher Scientific	S641
Dextrose	Ricca Chemical Company	RDCD0200
Dulbecco's PBS no Ca2+ or Mg2+	Thermo Scientific	SH3002802
Ficoll solution	GE Healthcare	17-1440-03
Acetic Acid	Fisher Scientific	BP2401
12 mm circular glass coverslips	Fisher Scientific	12-545-80 12CIR-1
24-well plates	Corning Incorporated	3524
Pooled Human Serum	Sigma	S7023
RPMI	Mediatech	15-040-CV
Fetal Bovine Serum	Thermo Scientific	SH3007103
Human interleukin-8	R&D Systems	208-IL/CF
MOPS	Sigma	M3183
MgCl2	Fisher Scientific	BP214
Propidium Iodide	Life Technologies	L7007 or L7012
SYTO9	Life Technologies	L7007 or L7012
Saponin	Fluka Analytical	47036
Alexa Fluor 647-coupled soybean lectin	Life Technologies	L-32463
DAPI	Sigma	D8417
SYTOX Green	Life Technologies	S7020
Mouse anti-CD63	Developmental Studies Hybridoma Bank	H5C6
Alexa Fluor 555 Antibody Labeling Kit	Life Technologies	A20187
Hemacytometer Bright Line	Hausser Scientific	1492
Forceps	EMS	78320
Sorvall Legend RT + Centrifuge	Thermo Scientific	75004377