Establishment and Comparison of Fluorescence-Based T6SS Activity Detection Methods in Acinetobacter baumannii

Summary

Here, we propose a quantitative detection method based on fluorescent labeling (especially Luciferase) to efficiently and accurately assess the activity of bacterial T6SS, which is suitable for high-throughput analysis of clinical strains.

Abstract

The Type VI Secretion System (T6SS) is a crucial mechanism mediating intercellular interactions in Gram-negative bacteria, particularly in pathogenic species such as Acinetobacter baumannii. Previous studies have shown that the large plasmid pAB3 in the A. baumannii ATCC 17978 strain encodes a TetR-like protein that inhibits the expression of core T6SS genes. In contrast, the WT^R- strain, which lacks pAB3, can stably express and secrete the T6SS effector protein Hcp and exhibits the ability to kill E. coli. The tssM gene, one of the core genes of T6SS, is essential for its activity; its deletion directly leads to the inactivation of T6SS. However, traditional T6SS activity detection methods, such as killing assays, suffer from low throughput and insufficient sensitivity. To address these limitations, we have developed quantitative detection methods based on fluorescent labeling.

To improve T6SS activity detection, we developed three fluorescent labeling methods: (1) A quantitative detection method based on Luciferase labeling, which is characterized by high specificity, sensitivity, and reproducibility, making it suitable for high-throughput analysis; (2) A detection method based on green fluorescent protein (GFP) labeling, which, despite being susceptible to environmental interference, offers the advantage of high throughput; (3) Flow cytometry detection, which can quantitatively assess bacterial viability but is operationally complex and costly. After a comprehensive comparison, the Luciferase-based labeling method proved to be the most accurate, sensitive, and user-friendly. When applied to 20 clinical isolates of A. baumannii, this method was confirmed to rapidly and accurately evaluate T6SS activity.

Introduction

The Type VI Secretion System (T6SS) is an important protein secretion system in Gram-negative bacteria, widely involved in bacterial competition, antagonism against eukaryotic hosts, and regulation of host immune responses^1,2. By injecting toxic effector proteins into neighboring bacteria or eukaryotic cells, T6SS helps bacteria maintain a competitive advantage in complex environments. In recent years, the significant role of T6SS in bacterial adaptability, pathogenicity, and microbial interactions has made it a research hotspot, especially in multidrug-resistant pathogens, where its functions provide potential targets for the development of new antibacterial strategies^3,4,5.

Acinetobacter baumannii (Ab) is a Gram-negative opportunistic pathogen widely distributed in hospital settings. Due to its high drug resistance and strong adaptability, it has become an important nosocomial pathogen worldwide^6,7,8. The survival of A. baumannii in complex microbial environments relies on multiple secretion systems, among which T6SS plays a key role in its competition with other microbes and interactions with the host. With its highly conserved T6SS gene cluster and well-defined functions, A. baumannii has become an ideal model for studying T6SS mechanisms. Moreover, the unique structure of the T6SS in A. baumannii provides a distinctive perspective for further revealing the diversity and functions of T6SS.

The T6SS gene cluster of A. baumannii contains 12 core protein genes (such as tssA-tssM, lacking tssJ) and several genes with unknown functions (such as tagX, tagN, tagF, etc.)^2,9,10. Among them, the hemolysin coregulated protein (Hcp) is an important component of T6SS, and its secretion is considered a hallmark of functional T6SS. Studies have shown that the large plasmid pAB3 in A. baumannii inhibits T6SS expression by encoding a TetR-like regulatory protein. In contrast, strains that have lost pAB3 (such as WT^R-) can stably express and secrete Hcp, exhibiting significant T6SS activity¹¹. The tssM gene is one of the core genes of T6SS, and its deletion directly leads to the inactivation of T6SS.

However, traditional T6SS activity detection methods (such as killing assays) have limitations such as low throughput, poor reproducibility, and insufficient sensitivity, which severely restrict the in-depth study and widespread application of T6SS⁵. Therefore, the development of efficient and accurate T6SS activity detection methods has become an urgent need in current research. This study aims to establish and compare T6SS activity detection methods based on fluorescent labeling to overcome the shortcomings of traditional methods. Through fluorescent labeling technology, we have achieved real-time, quantitative monitoring of T6SS activity, providing a more precise and reliable tool for the study of bacterial interactions. This study has validated the effectiveness of fluorescent labeling methods and, by comparing the advantages and disadvantages of different fluorescent labeling strategies, has provided new perspectives and methodological support for future research. It not only offers new technical means for T6SS research but also lays a scientific foundation for understanding the complex mechanisms of bacterial interactions and developing new antibacterial therapeutic strategies.

Protocol

1. Sources of strains and plasmids

1. Use A. baumannii standard strain ATCC 17978, E. coli (pZY01), and the clinical strain of A. baumannii isolated from the Department of Laboratory Medicine at the First Hospital of Jilin University.
2. Use the plasmid pZY01-GFP (carrying a kanamycin resistance gene, capable of stable expression in A. baumannii), which was modified and preserved in the laboratory, and the commercially purchased plasmid pGEN-luxCDABE (carrying an ampicillin resistance gene).
3. Culture A. baumannii and E. coli in Luria-Bertani (LB) medium or a minimal medium containing: 50 mM potassium phosphate buffer (pH 7.4), 15 mM (NH₄)₂SO₄, 2 mM MgSO₄, 0.2% (w/v) sodium succinate as the sole carbon source, and 0.01% (w/v) yeast extract¹². Add antibiotics at the following concentrations when necessary: E. coli: ampicillin (100 µg/mL) and kanamycin (50 µg/mL); A. baumannii: streptomycin (100 µg/mL).
   NOTE: E. coli (pZY01) and E. coli (pZY01-GFP) express kanamycin resistance. E. coli (pZY01) does not have a fluorescent marker so it can be referred to as E. coli in the following text.

2. Preparation of bacterial strains before the experiment

1. Retrieve the bacterial strains from the -80 °C freezer 2 days prior to the experiment. Inoculate the bacterial strains onto LB agar plates containing the corresponding antibiotics. Incubate the plates overnight at 37 °C.
   NOTE: Ensure that all cells retain the plasmids at the start of the experiment.
2. Pick two independent single colonies of each bacterial strain 1 day before the experiment. Inoculate each colony separately into two individual tubes containing 5 mL of LB liquid medium. Incubate the cultures overnight at 37 °C with shaking at 200 rpm. Use these cultures for subsequent experiments.
   NOTE: Optimize the incubation time based on the bacterial growth rate to ensure an adequate amount of bacterial biomass is obtained.

3. Preparation of E. coli (GFP)/ E. coli (Luciferase) strains

1. Transformation
   1. Add 50 ng of plasmid DNA (pZY01-GFP or pGEN-luxCDABE, quantified via NanoDrop at A260/A280) into 100 µL of E. coli DH5α competent cells, respectively, and mix gently.
   2. Incubate on ice for 30 min, heat-shock at 42 °C for 60 s, and immediately place on ice for 30 s.
   3. Add 700 µL of antibiotic-free LB broth and incubate at 37 °C with shaking at 180 rpm for 2 h.
2. Plating and culturing
   1. Centrifuge at 420 × g for 2 min, discard the supernatant, and resuspend the cell pellet in 100 µL of antibiotic-free LB broth.
   2. Spread the pZY01 (GFP) transformation product onto LB agar plates containing 50 µg/mL kanamycin.
   3. Spread the pGEN-luxCDABE transformation product onto LB agar plates containing 100 µg/mL ampicillin. Incubate at 37 °C overnight.
3. Verification of positive clones
   1. Pick single colonies and extract DNA as polymerase chain reaction (PCR) templates the next day. Perform PCR amplification and verify by agarose gel electrophoresis.
   2. Confirm positive strains and label them as E. coli (GFP) and E. coli (Luciferase).
      NOTE: Ensure the quality of competent cells is high for efficient transformation. Avoid excessive bacterial liquid during plating to prevent interference with single colony isolation.

4. Western blotting

1. Sample preparation
   1. Collect bacterial cultures and centrifuge (1500 × g, 10 min, 4 °C) to separate the cell pellet and supernatant.
   2. Add an appropriate amount of loading buffer to both the cell pellet and supernatant. Boil at 100 °C in a metal bath for 10 min. Centrifuge (1500 g, 10 min, 4 °C) and collect the supernatants as protein samples.
   3. Measure protein concentration using the bicinchoninic acid (BCA) or Bradford method and adjust to the same concentration.
2. Sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE)
   1. Prepare SDS-PAGE gels of appropriate concentrations (e.g., 12% separating gel, 5% stacking gel)¹³.
   2. Load 20 µg protein samples into the gel wells along with a 1-2 µL pre-stained protein marker. Perform electrophoresis at 100-120 V until the bromophenol blue indicator reaches the bottom of the gel.
      NOTE: Running Buffer: Tris 25 mM, Glycine 192 mM, SDS 0.1% (w/v), distilled water to 1 L.
3. Transfer (Western Blotting)
   1. Remove the gel from the glass plates and soak it in transfer buffer. Prepare a polyvinylidene difluoride (PVDF) or nitro cellulose (NC) membrane, activate the PVDF membrane with methanol, and then soak it in transfer buffer.
      ​NOTE: Transfer Buffer: 25 mM Tris, 192 mM Glycine, 20% Methanol
   2. Assemble the transfer apparatus in a "sandwich" structure (filter paper-gel-membrane-filter paper). Transfer at 100 V for 60-90 min (adjust time based on protein molecular weight; Host cell proteins (HCP) has a molecular weight of approximately 17 kDa).
      NOTE: Place the transfer tank in ice water and add ice packs to maintain a low temperature. If the transfer time is long, replace the ice packs midway.
4. Blocking
   1. After transfer, place the membrane in 5% skim milk-blocking solution. Block at room temperature (RT) on a shaker for 1 h or overnight at 4 °C.
5. Primary antibody incubation
   1. Wash the membrane three times with Tris-buffered saline (TBS) containing 0.1% Tween 20 (TBST) buffer, 5 min each time. Incubate the membrane with diluted primary antibodies (mouse anti-Hcp antibody, 1:1000 dilution; mouse anti-isocitrate dehydrogenase (ICDH) antibody, 1:1000 dilution) at 4 °C overnight or RT for 2 h.
6. Secondary antibody incubation
   1. Wash the membrane with TBST buffer 3 times, 5 min each. Incubate the membrane with diluted secondary antibody (Horseradish peroxidase [HRP]-conjugated anti-mouse IgG, 1:3000 dilution) at RT for 2 h.
7. Detection
   1. Wash the membrane with TBST buffer 3 times, 5 min each. Prepare ECL chemiluminescent substrate and evenly apply it to the membrane.
   2. Detect signals using a chemiluminescence imaging system and capture images (HCP protein has a molecular weight of approximately 17 kDa, ICDH is a cytoplasmic metabolic enzyme with an expected molecular weight of approximately 45 kDa in A. baumannii).
      NOTE: ICDH is an important quality control marker that can only be detected in the cell pellet fraction (containing intact bacteria) and not in the supernatant fraction (containing secreted proteins), thereby demonstrating that no cell lysis occurred during sample preparation. Hcp secretion (17 kDa) should only be visible in supernatants from T6SS-active strains.

5. Preparation of 96-well plates with LB agar medium

1. Quickly add autoclaved antibiotic-free LB agar medium, pre-warmed, into a sterile 96-well plate, 50 µL per well. Allow the plate to sit at RT for at least 1 h to ensure complete solidification of the medium.
2. Use black-walled plates with a transparent bottom for detecting GFP fluorescence.
3. Use light-proof 96-well plates for detecting Luciferase fluorescence to avoid cross-well fluorescence interference.
   NOTE: Avoid generating bubbles when adding the medium to ensure uniform gel thickness, which promotes optimal bacterial contact and growth.

6. A. baumannii killing assay

1. Standard killing assay
   1. Culture WT^R- and ΔtssM mutant strains overnight at 37 °C with shaking in LB medium supplemented with 100 µg/mL streptomycin, grow E. coli separately in LB medium containing 50 µg/mL kanamycin under identical conditions.
   2. Wash the bacterial cultures twice with sterile PBS and resuspend them in fresh antibiotic-free LB medium. Adjust the OD₆₀₀ to 1.0 for standardization.
   3. Mix WT^R- and ΔtssM mutant strains with E. coli at a 1:1 ratio.
   4. Spot 10 µL of the mixed bacterial suspension onto antibiotic-free LB agar plates and incubate at 37 °C for 6 h.
   5. Scrape the bacterial spots and resuspend them in 1 mL of PBS. Perform five 10-fold serial dilutions.
   6. Spot 10 µL of the diluted bacterial suspensions onto LB plates containing kanamycin and incubate overnight at 37 °C.
   7. Analyze the T6SS-dependent antibacterial activity of WT^R- and ΔtssM mutant strains by colony counting.
      NOTE: Ensure thorough mixing of bacterial suspensions to avoid experimental errors. Perform at least three independent replicates for each sample to ensure data reliability.
2. Quantitative detection of T6SS activity using luciferase labeling
   1. Standardize the OD₆₀₀ of E. coli (Luciferase), WT^R-, ΔtssM mutants, and other strains (e.g., ATCC 17978, E. coli, Ab#40, Ab#170, Ab#180) to 1.0. Add 5 µL of bacterial suspension to a 96-well plate containing antibiotic-free LB agar. After air-drying in the dark, measure luciferase-based bioluminescence to verify the luminescence specificity of E. coli (Luciferase).
   2. Perform a 5-fold serial dilution of E. coli (Luciferase). Spot 10 µL of the diluted bacterial suspension onto a 96-well plate with LB agar and an empty 96-well plate (both light-proof). Measure luciferase-based bioluminescence to evaluate the sensitivity of the luciferase-based detection method and confirm whether LB agar interferes with the luminescence signal.
   3. Mix E. coli (Luciferase) with WT^R-, ΔtssM mutants, and 20 clinical strains (OD₆₀₀ standardized to 1.0) at a 1:1 ratio. Dilute 10-fold, add 5 µL to a 96-well plate, and set up three technical replicates.
   4. After air-drying in the dark, measure bioluminescence. Incubate at 37 °C in the dark for 4-6 h, then measure bioluminescence again (emission wavelength: 562 nm).
   5. Quantitatively assess T6SS-dependent antibacterial activity by comparing bioluminescence values before and after co-culture.
      NOTE: Ensure thorough mixing of bacterial suspensions to avoid experimental errors. Perform at least three independent replicates for each sample to ensure data reliability. Use Student's t-test for statistical analysis.
3. Detection of T6SS activity using GFP labeling
   1. Follow the same procedure as the Luciferase labeling experiment (Excitation wavelength: 480 nm, emission wavelength: 509 nm).
   2. Mix E. coli (GFP) with WT^R- and ΔtssM mutants (OD₆₀₀ standardized to 1.0) at a 1:1 ratio. Spot 10 µL onto antibiotic-free LB agar plates. After air-drying at RT, capture images using the GFP fluorescence channel (Alex488 program, excitation: 488 nm, emission: 509 nm) of a multifunctional imaging system.
   3. Incubate at 37 °C in the dark for 4-6 h, then capture images again. Assess T6SS activity based on changes in fluorescence intensity of bacterial spots.
      NOTE: Perform all steps in the dark to avoid interference with fluorescence signals. Ensure thorough mixing of bacterial suspensions to avoid experimental errors. Perform at least three independent replicates for each sample to ensure data reliability.
4. Flow cytometry detection
   1. Mix WT^R- with E. coli (GFP) and ΔtssM mutants with E. coli (GFP) at a 1:1 ratio. Take an appropriate amount of the mixed bacterial suspension and dilute it to a suitable concentration (typically OD₆₀₀ of 0.1-0.5) using sterile PBS or 1× buffer.
      ​NOTE: 1× buffer refers to Flow Cytometry Staining Buffer (1×), the ingredients include: 1× PBS (137 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO₄, 1.8 mM KH₂PO₄, pH 7.4), supplemented with: 2-5% (v/v) heat-inactivated FBS (or 1% BSA as alternative), 2 mM EDTA.
   2. Centrifuge the diluted mixed bacterial suspension at 300 x g for 2 min and discard the supernatant. Resuspend the pellet in 1 mL of sterile PBS or 1× buffer and mix gently. Filter the resuspended solution through a 35 µm cell strainer to remove aggregates and ensure a single-cell suspension.
   3. Turn on the flow cytometer, preheat the laser, and calibrate the instrument.
      1. Set the detection parameters: use a 488 nm laser to excite GFP. Detection channel: FL1 (typically a 530/30 nm filter for GFP fluorescence detection). Set forward scatter (FSC) and side scatter (SSC) parameters to distinguish bacterial cells.
      2. Use unlabeled E. coli as a negative control, adjust the voltage and gain, and ensure the GFP-negative population is at the baseline of the fluorescence signal.
   4. Gently mix the filtered sample and transfer 100 µL to a flow cytometry tube.
      1. Analyze the samples using the following instrument settings: Set the FSC-H threshold at 19,000 to exclude debris, with gain values of 100 (FSC), 110 (SSC), and 80 (FITC).
      2. For gating, initially exclude debris and non-target signals in the FSC-A vs. SSC-A scatter plot, followed by population analysis in the FITC channel. Acquire a minimum of 10,000 events from the target population (defined by FSC/SSC characteristics) for each sample.
   5. Analyze the data using flow cytometry software (e.g., FlowJo, BD FACSDiva).
      1. Plot FSC vs. SSC scatter plots to gate the target bacterial population. Plot FL1 fluorescence histograms to distinguish GFP-positive and GFP-negative populations.
      2. Calculate the percentage of GFP-positive cells and the mean fluorescence intensity (MFI).
   6. Centrifuge the remaining mixed bacterial suspension at 300 g for 2 min and discard the supernatant. Resuspend the pellet in 10 µL of sterile water and spot it onto antibiotic-free LB agar plates.
   7. Incubate at 37 °C in the dark for 5 h, then scrape the bacterial spots and resuspend them in 500 µL of 1× PBS. Take 100 µL of the resuspended solution, dilute appropriately, and perform flow cytometry detection, repeating steps 6.4.4-6.4.5.
   8. Compare the percentage of GFP-positive cells and fluorescence intensity before and after co-culture. Evaluate the killing effect of WT^R- and ΔtssM mutants on E. coli (GFP).
      NOTE: Ensure a single-cell suspension to avoid cell aggregation affecting detection results. Calibrate the instrument before detection to ensure the accuracy and reproducibility of fluorescence signals. Avoid exposing samples to strong light during the experiment to prevent GFP fluorescence quenching. Perform at least three replicates for each sample to ensure data reliability.

Results

Traditional methods for detecting T6SS activity typically involve co-culturing predator and target strains on antibiotic-free LB agar plates for 5 h, followed by scraping the bacterial spots, resuspending them, and performing serial dilutions. The diluted bacterial suspension is then spotted onto selective agar plates. By counting the colonies of the target strain on the plates the next day, T6SS activity can be qualitatively assessed. However, this method has limitations, including cumbersome procedures, reliance on manual colony counting, and difficulty in quantitative analysis. To address these shortcomings, this study innovatively introduces a fluorescence-based quantitative detection method. Specifically, we labeled the target strain (e.g., E. coli) with fluorescent proteins (GFP or Luciferase) and mixed the predator and target strains at a 1:1 ratio followed by dilution. Before and after co-culture, the fluorescence values of the mixed bacterial suspension were measured, and T6SS activity was quantitatively assessed based on changes in fluorescence values, as shown in Figure 1. Compared to traditional methods, this fluorescence-based detection approach offers advantages such as simplified operation, quantitative results, and high reproducibility.

A. baumannii ATCC 17978 relies on T6SS to antagonize bacterial growth
A. baumannii ATCC 17978 carries a large plasmid named pAB3, which encodes two TetR-like regulatory proteins that suppress the expression of T6SS genes¹¹. Studies have shown that in the wild-type strain (WT) carrying the pAB3 plasmid, the expression and secretion of Hcp are significantly inhibited. In contrast, the pAB3-deficient strain, WT^R-, stably expresses and secretes Hcp, as shown in Figure 2A. To further investigate the function of T6SS, we constructed a ΔtssM mutant by knocking out the core component tssM of T6SS in the WT^R- strain. Experimental results revealed that while ΔtssM could synthesize Hcp normally, it completely lost the ability to secrete Hcp, indicating that the T6SS of ΔtssM is biologically inactive (Figure 2A). This result confirms the critical role of tssM in T6SS function.

This study further explored the killing ability of the WT^R- strain against E. coli mediated by T6SS. Using E. coli as the target strain and WT^R- and ΔtssM mutants as predator strains, with water as a blank control, predator and target strains were co-cultured at a 1:1 ratio for 5 h. The mixed suspension was then serially diluted and spotted onto selective solid media to evaluate the killing effect. The results demonstrated that the WT^R- strain exhibited significant killing activity against E. coli, while the ΔtssM mutant showed markedly reduced killing ability. This indicates that A. baumannii WT^R- can mediate bacterial killing through T6SS, as shown in Figure 2B.

Fluorescence-based detection of T6SS activity
Quantitative detection of T6SS activity using luciferase labeling
Before the experiment, we normalized the concentrations of the experimental strains and measured the bioluminescence values of equal volumes of E. coli, E. coli (Luciferase), WT^R-, Ab#40, Ab#170, and Ab#180. The results showed that only E. coli (Luciferase) produced detectable luciferase activity, as shown in Figure 3A, indicating that interference from other bacteria on bioluminescence is negligible and validating the specificity of this detection method. Subsequently, we performed serial dilutions of E. coli (Luciferase) (starting OD₆₀₀ = 1) and measured the bioluminescence values at different dilution factors. The results demonstrated that even after 7 rounds of 5-fold serial dilution, the bioluminescence values detected for E. coli (Luciferase) remained significantly higher than those of the blank control (p < 0.05), as shown in Figure 3B. Additionally, LB agar had almost no effect on the detection of luciferase-based bioluminescence, further enhancing the accuracy and reliability of this method, as shown in Figure 3B.

Based on the optimized conditions, we validated this method by conducting killing experiments using WT^R- and ΔtssM mutants as predator strains co-cultured with E. coli (Luciferase). The results showed that when E. coli (Luciferase) was cultured alone or co-cultured with the ΔtssM mutant for 5 h, its bioluminescence value increased significantly, indicating normal proliferation of E. coli (Luciferase) in the absence of T6SS-mediated killing. However, after 5 h of co-culture with WT^R-, the bioluminescence value of E. coli (Luciferase) decreased significantly (p < 0.05), as shown in Figure 3C, demonstrating the feasibility of the luciferase-based bioluminescence quantitative detection method in studying T6SS function.

Quantitative detection of T6SS activity using GFP labeling
We inserted the GFP (green fluorescent protein) gene into the plasmid pZY01 (kanamycin resistance), which can be stably expressed in E. coli, and transformed it into E. coli. This allowed us to distinguish the two bacterial strains under Alex488 excitation light, as shown in Supplementary Figure 1.

Before the experiment, to rule out potential interference from bacterial autofluorescence and ensure the accuracy of GFP-based quantitative detection of WT^R- T6SS-dependent antibacterial activity, we measured the fluorescence signals of GFP-labeled E. coli and unlabeled E. coli, as well as other strains. The results showed that GFP-labeled E. coli exhibited significant fluorescence signals, while unlabeled E. coli and other strains showed fluorescence levels comparable to background, as shown in Figure 4A. Subsequently, we validated the correlation between bacterial load and fluorescence intensity by measuring the fluorescence values of E. coli (GFP) at different dilution factors. The results demonstrated that even after 3 rounds of 5-fold serial dilution, the fluorescence intensity detected for E. coli (GFP) remained significantly higher than that of the blank control (p < 0.05), as shown in Figure 4B, confirming the quantitative reliability of this method.

In experiments using WT^R- and ΔtssM mutants as predator strains co-cultured with E. coli (GFP) to validate this method, we observed no significant change in fluorescence intensity after 5 h of co-culture between WT^R- and E. coli (GFP). This phenomenon may be attributed to two reasons: First, WT^R- effectively killed E. coli (GFP) through T6SS, preventing its further proliferation. However, GFP fluorescence does not depend on enzymatic or other cellular components, so even if E. coli (GFP) is killed, its fluorescence signal persists, leading to minimal changes in fluorescence intensity before and after co-culture. Second, the LB agar medium in the 96-well plates caused significant interference with fluorescence detection. As shown in Figure 4B, even blank wells (containing only LB agar) exhibited detectable fluorescence, and this background interference amplified detection errors, making it difficult to reflect minor changes in E. coli (GFP) load through fluorescence differences. In contrast, when E. coli (GFP) was cultured alone or co-cultured with the ΔtssM mutant for 5 h, the fluorescence intensity increased significantly, as shown in Figure 4C. This is because the T6SS function of the ΔtssM mutant is inactive, allowing E. coli (GFP) to proliferate rapidly during culture, resulting in a significant increase in fluorescence intensity (p < 0.05).

We also employed a plate titration method for detection. This method is based on the positive correlation between the fluorescence brightness of E. coli (GFP) colonies formed on antibiotic-free LB agar plates under the Alex488 program and the bacterial load. We mixed WT^R- and ΔtssM with E. coli (GFP) at a 1:1 ratio, spotted the mixture onto antibiotic-free LB plates, and co-cultured for 5 h. Fluorescence imaging was performed under the same exposure conditions. The results showed that the fluorescence brightness of colonies after co-culture of WT^R- and E. coli (GFP) was significantly lower than that of E. coli (GFP) cultured alone or co-cultured with the ΔtssM mutant, as shown in Figure 4D. This result visually reflects T6SS activity through changes in colony brightness.

In summary, the GFP-based fluorescence detection method and the plate titration method provide complementary technical approaches for studying T6SS activity. Although the fluorescence detection method offers significant advantages in terms of high throughput and convenience, it still suffers from background interference in quantitative analysis. On the other hand, the plate titration method provides qualitative validation of T6SS activity through changes in colony brightness.

Flow cytometry detection
GFP emits green fluorescence under the 488 nm excitation light of a flow cytometer¹⁴. Therefore, the GFP-positive population represents E. coli (GFP) stably expressing GFP, while the GFP-negative population represents WT^R- and ΔtssM mutants that do not express GFP. At the initial stage of the experiment, we mixed WT^R-, ΔtssM mutants, and E. coli (GFP) at a 1:1 ratio and analyzed the bacterial population composition using flow cytometry (Figure 5). The results showed that E. coli (GFP) accounted for approximately 50% of the total bacterial population before co-culture, as shown in Figure 5A, consistent with the experimental design.

After 5 h of co-culture, we again analyzed the changes in bacterial population composition using flow cytometry. The results revealed that after co-culture with WT^R-, the proportion of E. coli (GFP) significantly decreased (p < 0.05), as shown in Figure 6, indicating that WT^R- effectively inhibited or killed E. coli (GFP) through its functional T6SS. In contrast, after co-culture with the ΔtssM mutant, the proportion of E. coli (GFP) remained at approximately 50%, as shown in Figure 5B, indicating that the ΔtssM mutant, due to its inactive T6SS, could not inhibit or kill E. coli (GFP), allowing it to grow normally. These results align with expectations and further confirm the T6SS-dependent antibacterial activity of WT^R-.

In summary, flow cytometry provides an efficient and reliable method for quantitatively analyzing T6SS-dependent antibacterial activity by distinguishing GFP-positive and GFP-negative populations. However, flow cytometry indirectly proves bactericidal activity by comparing population proportions before and after co-culture. Additionally, its complex operation, high technical requirements, and significant costs make it unsuitable as the primary choice for large-scale screening.

Validation with clinical strains
Through a comprehensive evaluation of the aforementioned T6SS activity detection methods, the luciferase-based method demonstrated higher sensitivity compared to the GFP-based killing assay. As shown in Figure 3B, even after 7 rounds of 5-fold dilution, the bioluminescence values detected for E. coli (Luciferase) remained significantly higher than those of the blank control (p < 0.05), whereas the GFP-based method only achieved correlation up to 3 rounds of dilution. Additionally, the luciferase-based method was less affected by LB agar, further enhancing its accuracy and reliability (Figure 3B). The luciferase-based method, utilizing 96-well plates, offers high-throughput capabilities, enabling simultaneous detection of multiple samples and significantly improving experimental efficiency. Therefore, we concluded that the quantitative detection method based on Luciferase-labeled target strains exhibits significant advantages in accuracy, sensitivity, and operational simplicity. To validate the universality of this method, we further applied it to detect T6SS activity in 20 clinical isolates of A. baumannii. We used co-culture of WT^R- and E. coli (Luciferase) as a positive control and co-culture of ΔtssM mutant and E. coli (Luciferase) as a negative control.

First, we assessed the expression and secretion of Hcp in these clinical strains to preliminarily evaluate their T6SS functional status. In this study, Hcp secretion was considered indicative of T6SS activity under experimental conditions, but it does not universally equate to antibacterial activity, as T6SS function may depend on effector proteins, recipient cells, or environmental factors. The results showed that 10 strains-Ab#15, Ab#17, Ab#19, Ab#37, Ab#39, Ab#41, Ab#45, Ab#57, Ab#61, and Ab#63-were capable of expressing Hcp and secreting it into the supernatant, indicating that their T6SS was activated. In contrast, the remaining 10 strains, although capable of expressing Hcp, failed to secrete it into the supernatant, suggesting that their T6SS might be inactive or functionally impaired, as shown in Figure 6A.

Next, we used Luciferase-labeled E. coli as the target strain to conduct killing assays with the aforementioned 20 clinical strains. Before the experiment, all strains and E. coli (Luciferase) were adjusted to the same OD₆₀₀ value and mixed in equal volumes to ensure consistent initial conditions. The results showed that the initial fluorescence values of all experimental groups were at the same level, with no significant differences. After 5 h of co-culture, the bioluminescence values of the groups with strains Ab#15, Ab#17, Ab#19, Ab#37, Ab#39, Ab#41, Ab#45, Ab#57, Ab#61, and Ab#63 significantly decreased (p < 0.05), indicating that these strains exhibited strong killing activity against E. coli (Luciferase). This result was entirely consistent with the Hcp expression and secretion data from the Western blot experiments. Conversely, the remaining 10 strains that failed to secrete Hcp into the supernatant showed a significant increase in bioluminescence after co-culture with E. coli (Luciferase), indicating that these strains, due to inactive or impaired T6SS, were unable to effectively kill the target bacteria, as shown in Figure 6B.

In summary, the T6SS activity detection method based on Luciferase-labeled target strains not only offers high accuracy and sensitivity but also enables rapid, high-throughput assessment of T6SS activity in clinically isolated strains. This method provides robust technical support for systematic studies of T6SS function in clinical strains and opens new avenues for research into bacterial competition and pathogenic mechanisms.

Figure 1: Schematic diagram of conventional and novel T6SS killing experiments. In this experiment, E. coli was tagged with GFP or luciferase, and the killing ability of the attacker was quantified by measuring the fluorescence (for GFP) or luminescence (for luciferase) intensity produced by the prey E. coli cells after 5 h of co-culture with A. baumannii. Please click here to view a larger version of this figure.

Figure 2: A. baumannii ATCC 17978 relies on T6SS to antagonize bacterial growth. (A) The pAB3 plasmid represses the expression of T6SS in strain 17978. Culture supernatant of the specified bacterial strains was evaluated for secreted Hcp, one component of T6SS's puncturing structure. Note that the strain lacking pAB3 secretes Hcp and deletion of tssM eliminated such secretion. The cytosolic isocitrate dehydrogenase (ICDH) was used as an internal reference to assess the integrity of bacterial cells. (B) Cells of the wild-type A. baumannii or the ΔtssM mutant were mixed with the specified bacteria at a 1:1 ratio for 6 h. The survival of the prey bacteria was evaluated by plating dilutions on selective medium, the photo was taken under natural light (step 6.1). Please click here to view a larger version of this figure.

Figure 3: Quantitative detection of T6SS activity based on luciferase labeling. (A) Equal amounts of E. coli (luciferase) and other control bacteria were tested for luminescence values under the same experimental conditions to assess the specificity of the luciferase-labeled detection method (step 6.2.1). (B) After standardizing the OD₆₀₀ of E. coli (luciferase) to 1.0, the bacterial solution was diluted and spotted on a 96-well plate with transparent bottoms and light-shielded sides to measure luminescence values. These data were used to assess the sensitivity of the luciferase-labeled detection method and to confirm that LB Agar does not significantly interfere with the luminescence signal. (step 6.2.2). (C) Cells of WT^R- or ΔtssM mutants were mixed with E. coli (luciferase) at a 1:1 ratio and co-cultured for 5h. The change in luminescence values before and after co-culture was used to assess the WT^R- T6SS-dependent antibacterial activity. Statistical analysis was performed using Student's t-test, and the difference between WT^R- and ΔtssM groups was found to be significant (p < 0.05). Data represent the mean ± SD of three independent experiments (step 6.2.3-6.2.5). Please click here to view a larger version of this figure.

Figure 4: Quantitative detection of T6SS activity based on GFP labeling. (A) Equal amounts of E. coli (GFP) and other control bacteria were tested for fluorescence values under the same experimental conditions to assess the specificity of the GFP-labeled detection method. The bars represent the mean fluorescence values from three independent experiments, and error bars indicate the standard deviation (SD) (step 6.3.1). (B) After standardizing the OD₆₀₀ of E. coli (GFP) to 1.0, the bacterial solution was diluted and spotted on a 96-well plate with transparent bottoms and light-shielded sides to measure fluorescence values. These data were used to assess the sensitivity of the GFP-labeled detection method and to confirm that LB Agar does not significantly interfere with the fluorescence signal (step 6.3.1). (C) Cells of WT^R- or ΔtssM mutants were mixed with E. coli (GFP) at a 1:1 ratio and co-cultured for 5 h. The change in fluorescence values before and after co-culture was used to assess the WT^R- T6SS-dependent antibacterial activity. Statistical analysis was performed using Student's t-test, and the difference between WT^R- and ΔtssM groups was found to be significant (p < 0.05). Data represent the mean ± SD of three independent experiments (step 6.3.1). (D) Cells of WTR- or ΔtssM mutants were mixed with E. coli (GFP) at a 1:1 ratio and co-cultured for 5h. The change in brightness of plaques in bright-field imaging before and after co-culture was used to assess the WT^R- T6SS-dependent antibacterial activity (step 6.3.2-6.3.3). Please click here to view a larger version of this figure.

Figure 5: Detection of T6SS activity based on flow cytometry. (A) The proportion of E. coli (GFP) detected by flow cytometry before co-culture with WT^R- and ΔtssM. (B) The proportion of E. coli (GFP) detected by flow cytometry after co-culture with WT^R- and ΔtssM (protocol 6.4). Please click here to view a larger version of this figure.

Figure 6: Quantitative detection of the T6SS-dependent antibacterial activity of clinical strains by Luciferase-labeled E. coli. (A) The expression of Hcp in the pellet and supernatant of 20 clinical isolates was detected by immunoblotting. The presence of Hcp in the supernatant indicates that the T6SS of the strain is activated. The cytosolic isocitrate dehydrogenase (ICDH) was used as an internal reference to evaluate the integrity of bacterial cells. (B) The 20 clinical isolates of A. baumannii were co-cultured with E. coli (luciferase) at a 1:1 ratio for 5 h. The T6SS-dependent antibacterial activity of the clinical isolates of A. baumannii was assessed by the change in luminescence values before and after co-culture. Statistical analysis was performed using Student's t-test, and significant differences were observed between Hcp-secreting and non-secreting isolates (p < 0.05). Data represent the mean ± SD of three independent experiments (step 6.2.3). Please click here to view a larger version of this figure.

Supplementary Figure 1: E. coli (GFP) was imaged under a multimodal imaging system. Imaging of E. coli (GFP) cells on the same plate under GFP fluorescence (right) and its corresponding black-and-white version (left). Please click here to download this File.

Discussion

The Type VI Secretion System (T6SS) is a complex multi-protein machinery whose functions are primarily realized through the secretion of effector proteins. These effector proteins mediate interbacterial competition by killing or inhibiting other bacterial species, thereby providing the host bacterium with a competitive advantage in microbial communities¹⁵. Beyond its role in competition, studies have shown that T6SS is involved in various cellular processes, including bacterial colonization, environmental adaptation, biofilm formation, adhesion modification, and immune evasion¹⁶. Notably, the regulation of T6SS function is multi-layered, involving intricate mechanisms at the transcriptional, post-transcriptional, and post-translational levels¹⁷.

In the field of T6SS research, Acinetobacter calcoaceticus and Acinetobacter baumannii have emerged as model strains due to their unique advantages. First, the genome of A. baumannii has been extensively sequenced and annotated, providing researchers with a wealth of genetic information¹⁸. Second, this strain possesses a highly functional T6SS, with key components such as Hcp having been thoroughly characterized in terms of structure and function¹². Research has demonstrated that T6SS not only participates in bacterial competition but is also closely related to the virulence of A. baumannii, playing a critical role in bacterial pathogenesis¹⁹. In recent years, significant progress has been made in T6SS research in A. baumannii, particularly in the functional characterization of key effector proteins (e.g., VgrG, PAAR, Hcp) and effector-immunity pairs². Additionally, breakthroughs have been achieved in understanding the regulatory network of T6SS, with transcription factors such as integration host factor (IHF) and HapR identified as regulators of T6SS expression²⁰.

With the continuous discovery of T6SS effectors, related research methods are also evolving. The application of quantitative transcriptomics and proteomics technologies has enabled researchers to comprehensively assess the impact of T6SS effectors on bacterial physiology²¹. By constructing T6SS-deficient and complemented strains, researchers can precisely dissect the specific mechanisms of T6SS in bacterial competition and pathogenesis²². However, as novel T6SS effectors and regulatory factors continue to be discovered, existing research methods still face limitations in sensitivity and throughput, highlighting the need for more advanced research tools.

This study successfully established and optimized a fluorescence-based T6SS activity detection method, effectively overcoming the limitations of traditional approaches. By employing fluorescence labeling technology, we achieved real-time, quantitative monitoring of T6SS activity, providing a more precise and reliable technical platform for studying bacterial interactions. The results demonstrated that, compared to traditional killing assays, the Luciferase-based detection method offers significant advantages: first, it exhibits higher sensitivity and accuracy, enabling quantitative detection of T6SS-dependent antibacterial activity; second, its high-throughput capability significantly enhances research efficiency, facilitating large-scale screening of T6SS inhibitors or activators. Notably, the Luciferase detection method demonstrates excellent reproducibility, with stable luminescence signals that are less susceptible to variations in experimental conditions, primarily due to its unique biochemical luminescence mechanism²³.

The bacterial luciferase system (luxCDABE) represents a fundamentally distinct mechanism from eukaryotic luciferases, characterized by an enzyme complex that catalyzes light emission through FMNH₂-dependent oxidation of long-chain aldehydes²⁴. In our engineered E. coli (luxCDABE) system, three critical factors govern detection sensitivity and reliability: (1) The endogenous aldehydes produced by LuxCDE enzymes exhibit nonlinear accumulation kinetics during bacterial growth²⁵; (2) Strict oxygen dependence limits signal detection in hypoxic microenvironments²⁶; and (3) Temperature sensitivity, with optimal activity observed at 30-37 °C (rapid inactivation occurs above 40 °C)²⁷. Our optimization experiments demonstrated that maintaining cultures at 30 ± 0.5 °C in pH-buffered medium (7.8 Tris-HCl) achieved maximum signal stability. Although environmental factors such as temperature fluctuations could cause signal variations in our tests, normalization to OD₆₀₀-controlled cell density effectively compensated for these effects. This method enables real-time monitoring of T6SS activity without the need for exogenous substrate addition. However, researchers should consider the inherent limitations of this system: Signal reduction in high-density cultures due to oxygen depletion²⁸, and Early-phase nonlinear signals during aldehyde precursor accumulation²⁵. These biochemical constraints highlight the importance of standardized culture conditions when employing bacterial luciferase as a quantitative tool for studying interbacterial competition.

In contrast, the GFP-based detection method has notable limitations. The fluorescence properties of GFP require light-protected operations throughout the experiment, and its fluorescence intensity decays over time, leading to poor reproducibility²⁹. Additionally, variations in the thickness of agar medium in 96-well plates can significantly affect fluorescence detection results, particularly when dealing with strains with weak killing activity, potentially introducing substantial errors. However, flow cytometry can visually reflect T6SS-dependent antibacterial activity, its complex operation and low throughput limit its application.

Based on the findings of this study, we recommend the following optimizations for future research: first, further refine detection conditions and explore the potential of this method in studying other bacterial secretion systems; second, integrate the Luciferase detection method with high-throughput technologies such as transcriptomics and proteomics to comprehensively dissect the complex regulatory network of T6SS; and finally, provide new theoretical foundations and therapeutic strategies for the prevention and treatment of bacterial diseases through in-depth research on the functional mechanisms of T6SS.

The Luciferase-based T6SS activity detection method developed in this study offers significant advantages in sensitivity, accuracy, and throughput, providing robust technical support for T6SS functional research. This method not only aids in elucidating the mechanisms of T6SS in bacterial pathogenicity, symbiosis, and competition but also facilitates the identification and functional characterization of novel T6SS effectors. However, researchers must strictly control experimental conditions and consider the method's applicability and limitations. Future research should focus on optimizing detection conditions and expanding its application to the study of other bacterial secretion systems, which will contribute to a comprehensive understanding of the functional characteristics and regulatory mechanisms of T6SS.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
10%, 15% SDS-PAGE Gel Preparation Kit	Epizyme	PG112
-20 °C Freezer	Haier	HYCD-290
37 °C Incubator	Bluepard	181254254
4 °C Refrigerator	Haier	HYC390
-80 °C Freezer	Haier	DW-86L726G(726L)
96-well plate	JET	TCP010096
Agar	Biofrox	8211KG001
Ammonium Persulfate (APS)	Thermo	7727-54-0
Biological Safety Cabinet	ESCO	AC2-4S1
Electronic Balance	Sartorius AG	BSA124S-CW
Electrophoresis apparatus	BIO-RAD	POWER PAC1000
Flake Ice Machine	GRANT	XB70
Flow Cytometer	BECKMAN COULTER	AW38143
Flow Cytometry Staining Buffer	proteintech	PF00018
Gel Imaging System	BIO-RAD	Gel Doc 2000
Glycine	Zike	ZK-L2577
High-speed Centrifuge	Eppendorf	5405IN106358
HRP-conjugated Rabbit/Mouse Secondary Antibody	proteintech	SA00001-2
LB Broth	Solarbio	L8291
Low Temperature High Speed Centrifuge	Thermo	17R
Methanol	Thermo	R40121
Micro UV-Vis Spectrophotometer	Thermo	Nanodrop one
Microcentrifuge	allsheng	Mini-6k
Microplate Reader	Bio-Tek	H1M
NuPAGE LDS Sample Buffer	Thermo	NP0007
Phosphate Buffer Solution (PBS)	Zike	ZK-L1649
Precision Plus Protein Dual Color Standards	Bio-Rad	1610374
SDS Sample Loading Buffer	Beyotime	poo15L
Skim Milk Powder	Thermo	LP0033B
Sodium Dodecyl Sulfate (SDS)	Zike	zk6885
Thermostatic Water Bath	JingHong	DK-420S
Transfer Apparatus	Bio-Rad	PowerPac HC
Tris(hydroxymethyl)aminomethane (Tris)	Zike	ZK-L2557
Vertical Electrophoresis Apparatus	Bio-Rad	Mii-PROTEAN Tetra