Monitoring single cell bioenergetic status and cell lysis in dense and differentiating Bacillus subtilis cultures

Monitoring single cell bioenergetic status and cell lysis in dense and differentiating Bacillus subtilis cultures | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Monitoring single cell bioenergetic status and cell lysis in dense and differentiating Bacillus subtilis cultures Maria Dakes Stavrakakis , View ORCID Profile Madeleine Humphrey , View ORCID Profile Tjeerd van Rij , View ORCID Profile Colin R. Harwood , View ORCID Profile Henrik Strahl doi: https://doi.org/10.1101/2025.08.21.671461 Maria Dakes Stavrakakis 1 Centre for Bacterial Cell Biology, Biosciences Institute, Newcastle University , Richardson Road, Newcastle upon Tyne, NE2 4AX, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site Madeleine Humphrey 1 Centre for Bacterial Cell Biology, Biosciences Institute, Newcastle University , Richardson Road, Newcastle upon Tyne, NE2 4AX, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Madeleine Humphrey Tjeerd van Rij 2 DSM Biotechnology Center , Alexander Fleminglaan 1, Delft 2613 AX, Netherlands Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Tjeerd van Rij Colin R. Harwood 1 Centre for Bacterial Cell Biology, Biosciences Institute, Newcastle University , Richardson Road, Newcastle upon Tyne, NE2 4AX, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Colin R. Harwood For correspondence: colin.harwood{at}newcastle.ac.uk h.strahl{at}newcastle.ac.uk Henrik Strahl 1 Centre for Bacterial Cell Biology, Biosciences Institute, Newcastle University , Richardson Road, Newcastle upon Tyne, NE2 4AX, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Henrik Strahl For correspondence: colin.harwood{at}newcastle.ac.uk h.strahl{at}newcastle.ac.uk Abstract Full Text Info/History Metrics Supplementary material Data/Code Preview PDF ABSTRACT Bacillus subtilis is a major model organism for studying population heterogeneity in clonal bacterial cultures due to its high genetic tractability and ability to differentiate into subpopulations with distinct biological functions. It is also a key industrial production host, responsible for synthesizing a range of commercially valuable enzymes and metabolites. However, cell differentiation processes can pose a challenge for the optimal biotechnological utilization of B. subtilis , particularly when emerging subpopulations do not contribute to product biosynthesis. Here, we present robust assays that facilitate the analysis of two previously difficult-to-study population properties of B. subtilis : (i) the energization levels of individual cells within dense cultures and (ii) the extent of cell lysis that can occur under such conditions. Our findings reveal an unappreciated level of heterogeneity in cell energization within dense B. subtilis cultures, and a surprisingly high degree of cell lysis in seemingly healthy, actively growing populations. These insights add to our understanding of the biological complexities and single-cell heterogeneities present in superficially simple bacterial clonal cultures, establish analytical tools to study the associated processes, and provide a foundation for further optimizing B. subtilis as an industrial production host. IMPORTANCE Bacillus subtilis and its close relatives are important industrial microorganisms, responsible for the production of a range of commercially valuable enzymes, antibiotics and metabolites. In recent years, considerable research efforts have been aimed at increasing the productivity of these organisms. However, their ability to undergo physiological and morphological differentiation processes at high cell densities ultimately limits their productivity. Our research reveals how the resulting heterogeneity impacts the population-level energy status of individual cells in the culture and the surprisingly high extent of population-level cell lysis. It also provides tools for determining these important productivity criteria as well as guiding the development of the production host. INTRODUCTION Although bacteria are often described as free-living single-celled organisms, in their natural environment, they participate in a range of social behaviours that influence their long-term survival. In many cases, this involves the formation of complex communities, with both kith and kin, that can either be cooperative (beneficial) or antagonistic (competitive) ( 1 ). The Gram-positive endospore-forming bacterium Bacillus subtilis has been shown to form such complex communities in which bet-hedging and social differentiation are important survival strategies ( 2 , 3 ). Ultimately, this involves the morphological differentiation (sporulation) of a minority of the cell population into dormant and highly temperature, desiccation, and chemically resistant endospores ( 4 ). However, the formation of spores is a complex and energy-demanding process that requires significant remodelling of metabolic activities and cellular structures which, furthermore, must take place in a nutritionally challenging environment. As a result, B. subtilis communities strive to delay the commitment to the sporulation differentiation pathway by seeking and utilising alternative nutrient sources ( 5 ). This is achieved by developing clonal, yet physiologically distinct cell types specialised in foraging (motility, swarming and chemotaxis), horizontal gene transfer (competence and transformation), utilisation of complex substrates (secretion of hydrolases), and production of antagonists to combat competitors (secondary metabolites and antimicrobial peptides) ( 5 – 8 ). This ability to differentiate into subpopulations with distinct phenotypes provides B. subtilis with a selective advantage in its native environment, the soil. In addition to the above-mentioned, well-defined differentiation pathways, Bacillus subtilis exhibits autolytic behaviour upon de-energisation triggered by extreme nutrient-limiting conditions ( 9 , 10 ). Thereby, a proportion of the cell population undergoes cell lysis, which releases nutrients allowing the remaining cell population to sustain metabolic activity and energisation ( 11 ). This allows the population to delay or even avoid the induction of the energy-intensive differentiation pathway leading ultimately to sporulation. In addition to relying on stochastic processes leading to deenergisation and the associated autolysis of a sub-population, cells in the early and still reversible stages of sporulation synthesise and secrete two proteins, the sporulation delaying protein (SdpC) and the sporulation killing factor (SkfA) that trigger lysis of other members of the community ( 12 ). This represents a form of active cannibalism that affects a proportion of the non-sporulating cells, ultimately providing an additional source of nutrients for the remaining cell population ( 13 ). In addition to serving as a model for the study of Gram-positive bacteria, B. subtilis and its close relatives are important industrial organisms, responsible for producing a range of industrial enzymes, peptide antibiotics, and metabolites ( 14 ). While the complex differentiation processes and the associated population heterogeneities provide an advantage in the natural environment, they can represent a distinct disadvantage in industrial fed-batch bioreactor cultivation conditions, where optical densities of over 100 are commonly required ( 15 ). In particular, both programmed and energy depletion-linked autolysis can reduce productivity and release into the culture medium intracellular products, such as nucleic acids and proteases, that can be detrimental to the final product ( 16 ). This means that attempts to reduce cell lysis by engineering production strains and optimising fermentation conditions require the monitoring of both the energetic status of individual cells, and the extent of cell lysis at different stages of growth. At the single cell level, bacterial viability and lysis is commonly monitored by so-called viability or Live/Dead assays ( 17 ), which find their use in a wide range of applications including microbiological quality control assessments of environmental or industrial samples and studies on the effectiveness and mode of action of antibiotics ( 18 ). Despite the arguably misleading commercial names, these types of assays do not report viability per se but utilise membrane-impermeable DNA-intercalating fluorescent dyes such as SYTOX Green and propidium iodide, which are excluded from bacterial cells with intact cell membranes ( 19 , 20 ). When the membrane permeability barrier is compromised through membrane-active compounds or when the cells undergo cell lysis, these dyes are able to penetrate the cell and stain cellular DNA ( 21 , 22 ). However, these assays cannot detect more subtle disturbances of the membrane permeability barrier function such as smaller-sized pores or increased ion conductivity and, indeed, are blind to metabolic inactivity unless associated with a severe membrane disruption or cell lysis. In contrast, voltage-sensitive fluorescent probes such as the carbocyanine dye, 3,3’-dipropylthiadicarbocyanine iodide (DiSC 3 ( 5 )) can be used to assay cell membrane potential, which provides a more direct readout for a cell’s bioenergetic status and a more sensitive proxy for the state of membrane permeability barrier function. Because of its cationic and hydrophobic characteristics, DiSC 3 ( 5 ) accumulates in polarised cells in a voltage-dependent manner. Upon membrane depolarisation, the dye is released from the cell, resulting in a lower fluorescence signal. This voltage-dependent staining behaviour allows the membrane potential levels to be monitored at the single-cell level using flow cytometry or fluorescence microscopy ( 23 , 24 ). The use of DiSC 3 ( 5 ) as a single-cell reporter in B. subtilis is well established ( 23 , 25 ). However, due to cell density-related complexities in staining and the maintenance of native membrane energisation during sample processing, the use of DiSC 3 ( 5 ) has been limited to early logarithmic growth phase or low cell density cultures, restricting comparisons to cultures with similar optical densities. Consequently, there is a lack of reliable protocols for monitoring bacterial bioenergetic status using voltage-sensitive dyes at the single-cell level from more varied culture conditions. In this manuscript, we provide a systematic analysis of factors relevant to measuring B. subtilis membrane potential across a range of cell densities, thereby significantly expanding the applicability of DiSC 3 ( 5 ) as a sensitive proxy for cellular bioenergetic status. Furthermore, we establish how a combination of DiSC 3 ( 5 ) and SYTOX Green can be used to monitor cell energisation levels and cell lysis across the different growth phases of B. subtilis . This method provides insights into an unexpectedly high level of population heterogeneity in cell energisation and lysis that can occur in a well-growing culture. While our study is limited to B. subtilis , we are confident that the established methods provide a solid foundation for developing similar assays in other microorganisms. MATERIALS AND METHODS Strains, media and growth conditions The used bacterial strains and associated genotypes are listed in Table 1 . Strain 168 ( trpC2 ) is a tryptophan auxotroph due to a three-base pair deletion in trpC , encoding indol-3-glycerol-phosphate synthetase (Fig. S1). We isolated a prototrophic variant of 168 to avoid the need to add tryptophan to the minimal salts culture medium. Previous attempts to generate a prototrophic strains of 168 often used DNA from strain W23, which is only 94.6% identical to 168 ( 26 ). To ensure the restoration of an authentic prototrophic version of trpC , we transformed strain 168 with a PCR-generated product of the trpC gene from strain NCIB3610 T , amplified using oligos 5’-CTTCACAACCTCACTCCCTAAACAAAGC and 5’-GGAAGAGATCTATGCTTGAAAAAATCATCAAAC. The trpC gene of NCIB3610 T is identical to that of 168 except for the absence of a 3-bp deletion at nucleotides 423-5 ( 26 ). Following selection on minimal medium plates, transformant MDS23 was purified and genome sequenced to confirm this strain carries the prototrophic trpC gene, while being otherwise genetically identical to the reference strain 168 ( 27 ) (Fig. S1). View this table: View inline View popup Download powerpoint Table 1: The list of bacterial strains used in this study. The list describes the name, genotype and origin of the strains. Antibiotic resistance markers: tetracycline ( tet ), spectinomycin ( spec ), neomycin ( neo ), kanamycin ( kan ), chloramphenicol ( cat ). Overnight cultures of B. subtilis were routinely grown in Lysogeny broth medium (LB; 0.5% (w/v) yeast extract, 1% (w/v) tryptone, 1% (w/v) NaCl), at 30°C and supplemented with 0.2% glucose to suppress sporulation. For the main experiments, strains were diluted 1:100 and grown at 37°C in modified Spizizen-minimal medium (mSMM: 0.2% (w/v) (NH 4 ) 2 SO 4 , 0.6% (v/v) KH 2 PO 4 , 1.4% (v/v) K 2 HPO 4 , 0.2% (v/v) K + -glutamate, 0.12% (v/v) trisodium citrate (Na 3 C 6 H 5 O 7 × 2H 2 O), 0.02% (v/v) MgSO 4 × 7H 2 O, 0.005% (v/v) ammonium ferric citrate ((NH 4 ) 5 Fe(C 6 H 4 O 7 ) 2 ), 0.5% (v/v) glucose, 0.02% (v/v) casamino acids). When growing the tryptophan auxotrophic B. subtilis strain 168, 0.002% (v/v) L-tryptophan was also added to the minimal medium. The autolysis experiments shown in Figure 1 and Supplementary Movie 1 were carried out in LB medium. Download figure Open in new tab Figure 1: Autolysis of B. subtilis can be monitored with the membrane permeability-indicator SYTOX Green (a) B. subtilis phase contrast and fluorescence time-lapse microscopy images of cells stained with 200 nM SYTOX Green in the presence of 100 μM of the proton uncoupler CCCP. For the full time-lapse experiment, see Movie S1. (b) B. subtilis phase contrast and fluorescence microscopy images of cells stained with 200 nM SYTOX Green in the absence (left panels) and presence (right panels) of the membrane-targeting, pore-forming lantibiotic nisin (10 μM for 5 min). The images are presented using identical contrast settings allowing a direct comparison between phase dark/SYTOX negative cells with intact membranes (left panels), phase dark/SYTOX Green positive cells with permeabilised membranes (middle panels) and extensively lysed phase light/ SYTOX Green positive cells (right panels). Scale bar: 3 µm. Strain used: (a) B. subtilis wild type (168) and (b) B. subtilis prototroph (MDS23). Fluorescence microscopy For fluorescence microscopy, the cells were incubated with either 1 µM of the voltage-sensitive dye DiSC 3 ( 5 ) dissolved in DMSO (Sigma-Aldrich) or 200 nM of the membrane impermeable fluorescent dye SYTOX Green dissolved in H 2 O (Thermo Fisher) for 5 min prior to imaging. 1% DMSO was maintained upon staining with DiSC 3 ( 5 ) to aid dye solubility. For the determination of the membrane potential using DiSC 3 ( 5 ), the cells were diluted to an OD 600 of 0.3 using spent medium obtained by centrifugation of an aliquot from the same culture. As positive controls for membrane depolarisation or membrane permeabilisation, the cells were incubated with either 10 μM of gramicidin or 10 μM nisin for 5 min prior to imaging. Incubations with dyes or antibiotics were performed by transferring 200 μl of cells into a 2 ml microcentrifuge tube with a perforated lid (to allow aeration), and shaking at 850 rpm and 37°C in a ThermoMixer (Eppendorf) for 5 min. For visualisation, the cell suspensions were immobilised on Teflon-coated multi-spot microscope slides (Hendley-Essex) covered by a thin layer of 1.2% (w/v) agarose. For membrane potential measurements, the slide was pre-warmed to 37°C prior to imaging. A sample of the cell suspension (0.5 µl) was applied to the agarose surface, air-dried, and covered with a coverslip. The slides prepared in this manner were imaged within 5 min of the application of the coverslip to minimise oxygen limitation and the associated loss of membrane potential. See Winkel et al. ( 23 ) for further details regarding slide preparation and time-lapse microscopy. The phase contrast and fluorescence imaging were carried out using a Nikon Eclipse Ti microscope (Nikon Plan Apo 100x/1.40 Oil Ph3 objective) equipped with Semrock Cy5-4040C (EX 628/40, DM660lp, EM 692/40) and Chroma 49002 (EX470/40, DM495lpxr, EM525/50) filter sets for imaging DiSC 3 ( 5 ) and SYTOX Green, respectively. The images were acquired using Metamorph 7.7 (Molecular Devices, Inc). Microscopy analysis For fluorescent microscopy, images were analysed using the Fiji software package ( 19 , 20 ). Quantification of DiSC 3 ( 5 ) fluorescence was performed in a semiautomated manner. In brief, the individual cells were identified as regions of interest (ROI) by thresholding the phase contrast images. If cells grew as clusters that could be visually identified as individual cells but that could not be separated based on the image thresholding, the ROIs were separated by a manually drawn thin line. The respective fluorescence images were background-subtracted prior to analysis. The mean fluorescence intensity values for single cells were acquired using a custom script available at https://github.com/NCL-ImageAnalysis/General_Fiji_Macros . Fluorometric measurement of lysis using SYTOX Green For the determination and quantification of cell lysis in a B. subtilis culture during growth for 15 h, culture supernatant containing the lysis-derived DNA was collected by centrifuging 500 μl of culture in a 1.5 ml Eppendorf tube (5 min, RT, 16,000 x g ). The supernatant was diluted 1:10 in a black, polystyrene, flat-bottomed microtiter plate (Porvair Sciences), and 1 μM of the DNA-intercalating dye SYTOX Green (Thermo Fisher Scientific) was added and incubated while shaking at 200 rpm at room temperature in a BMG Clariostar multimode plate reader using 485-10 nm excitation and 520-10 nm emission wavelength windows, respectively. Fluorescence intensity was monitored until stable levels were obtained. The background fluorescence of SYTOX Green in medium-only samples was subtracted from the test samples. For the preparation of the SYTOX Green-based lysis, a calibration curve was prepared from samples of B. subtilis grown as described above to different cell densities. Culture samples (500 μl) were collected at various optical densities and sonicated on ice for 10 min to lyse the cells. The supernatants of the sonicated samples were subjected to the fluorometric SYTOX Green assay as described above. The resulting calibration curve was used to convert the measured SYTOX Green signal to OD 600 of the cell population pre-lysis, assuming the full release of the cellular DNA content into the culture supernatant. The percentage of cell lysis was then calculated as follows: Percentage cell lysis = (calculated pre-lysis OD 600 of the lysing cell population)/(OD 600 of culture)*100. Statistical analysis Statistical analysis was performed using GraphPad Prism 9.5, making use of either an ordinary one-way, unpaired ANOVA with a Dunnett’s multiple comparisons test or an unpaired, two-sided t-test. RESULTS AND DISCUSSION Use of membrane permeability indicator to monitor autolysis in a B. subtilis culture If the majority of a non-growing bacterial cell culture undergoes lysis, this can be straightforwardly monitored as a decline in culture optical density. However, if the lysis process only affects a subpopulation of a growing culture, detecting lysis is less trivial and the associated reduced rate of optical density increase can easily be misinterpreted as a reduction in growth rate. An alternative to monitoring changes in culture optical density is the use of so-called viability indicators or Live/Dead stains such as SYTOX Green or propidium iodide, which detect the presence of cells with compromised membrane integrity ( 19 , 20 ). These assays rely on DNA-intercalating dyes that exhibit strongly enhanced fluorescence upon DNA binding but are membrane impermeable and thus unable to stain the DNA of cells with intact cell membranes. While these dyes are more commonly used to detect the activity of directly membrane-disrupting compounds ( 25 , 29 ), the integrity of a bacterial cytoplasmic membrane is disrupted as part of the autolytic process, thus potentially allowing detection of autolysis as well. However, to facilitate a robust interpretation of lysis data, it is crucial to consider the fate of cytoplasmic DNA during the autolytic process. CCCP is an ionophore that transports protons across the bacterial cytoplasmic membrane, thus triggering a collapse of the proton motive force (pfm) and an associated decline in cell ATP levels in B. subtilis ( 9 ). This comprehensive de-energisation induces misregulation of cell wall hydrolases, ultimately leading to induced autolysis ( 30 , 31 ). To monitor the fate of cellular DNA following B. subtilis autolysis, we followed CCCP-induced lysis using time-lapse microscopy and SYTOX Green staining. As shown in Figure 1a and Movie S1, autolysis indeed triggers permeabilisation of the cytoplasmic membrane that can be detected with SYTOX Green prior to significant changes in phase contrast becoming evident. However, the fate of the cellular DNA is rather heterogeneous with both rapid release to the medium and retention within the lysed cell remnants being observed. Similar heterogeneity is evident when autolysis is induced with the pore-forming antimicrobial peptide Nisin ( Figure 1b ), which forms large pores in B. subtilis membranes in a lipid II-dependent manner ( 32 , 33 ). While the pore-forming activity allows SYTOX Green to rapidly enter the cell and stain the cytoplasmic DNA, extended incubation that leads to autolysis is associated with at least partial release of the cellular DNA. In conclusion, while SYTOX Green fluorescence can be used to monitor the autolytic process, the associated release of cellular DNA complicates the analysis at the single-cell level unless time-lapse microscopy is applied. Nonetheless, SYTOX Green does gain access to DNA upon autolysis, which should enable the development of a population-based bulk assay. Analysing autolysis with a membrane permeability indicator in culture-based bulk assays SYTOX Green and Propidium Iodide (PI) are frequently used in fluorescence plate reader-based bulk assays to measure changes in membrane permeability as a proxy for cell viability ( 19 , 20 , 32 ). However, these assays are typically used to monitor a relatively rapid response to compounds and conditions that directly affect membrane-permeabilising properties. When we monitored SYTOX Green and PI fluorescence in B. subtilis cultures grown into the late stationary growth phase using a fluorescence plate reader, no clear signature for autolysis could be detected even though we expected the stationary, nutrient-starved cultures to exhibit significant levels of autolysis (Supplementary Figure 2). Thus, autolysis affecting part of the B. subtilis cell population cannot be easily detected by monitoring changes in SYTOX Green and PI fluorescence of the culture. To confirm that stationary growth phase B. subtilis cultures indeed undergo partial autolysis, we determined whether DNA released upon lysis can be detected in culture supernatants during early stationary phase ( Figure 2a ). This was achieved by staining a sample of the culture with SYTOX Green and assaying the SYTOX Green fluorescence signal. While there was a strong fluorescence signal in the culture supernatant fraction, only a weak signal was detected in the pellet fraction ( Figure 2b ). This suggests that a significant degree of autolysis is indeed occurring in such a B. subtilis culture, and that most of the DNA of lysed cells ultimately accumulates in the culture supernatant rather than remaining trapped within lysed cells. Finally, the presence of DNA in the culture supernatant was further confirmed by running the sample on an agarose gel stained with the DNA dye Nancy-520 ( Figure 2c ). Download figure Open in new tab Figure 2: Extracellular DNA is present in the stationary growth phase B. subtilis cultures ( a ) Growth kinetics of B. subtilis grown in mSMM at 37°C with the black arrow indicating the time point of sample collection. (b) SYTOX Green fluorescence signal measured fluorometrically for the cell pellet and the corresponding culture supernatant. The graph depicts the mean and standard deviation from three independent biological replicates. (c) Agarose gel depicting DNA stained with Nancy-520 in one of the supernatant samples analysed in panel (b). (d) The SYTOX Green fluorescence signal for an undigested culture supernatant sample, and for samples digested with 10 Kunitz units of bovine pancreas DNase I for 1 h at 37°C in the presence and absence of a supplemented divalent cation (20 mM MgSO 4 ). The graph depicts the mean and standard deviation from three independent biological replicates, along with P values of a one-way, unpaired ANOVA. (e) Agarose gel depicting DNA stained with Nancy-520 in the same supernatant samples as shown in panel (d) . The 1 kb DNA Ladder from NEB® was used as a molecular size marker. Strain used: B. subtilis prototroph (MDS23). **** = p ≤ 0.0001; ns = no significant difference. To verify that the SYTOX Green signal detected in the culture supernatant was due to the presence of DNA, the supernatant samples were incubated with endoribonuclease DNase I in the presence or absence of supplementary Mg 2+ , an essential cofactor for activity ( 33 , 34 ). When supernatant samples were incubated with DNase I in the absence of supplementary Mg 2+ , a fainter smear of lower molecular sized DNA was observed. However, no significant reduction of the corresponding SYTOX Green signal was observed ( Figure 2d, e ). In contrast, when supplementary Mg 2+ , was added to the supernatant along with DNase I, the SYTOX Green signal was significantly reduced and no DNA was detected in the agarose gel, indicating extensive DNA degradation ( Figure 2d, e ). Thus, the signal detected by SYTOX Green in culture supernatant is specific for DNA. Endogenous nucleases can potentially degrade DNA released when cells lyse. B. subtilis encodes two major and one minor extracellular nuclease: the membrane-associated nuclease NucA, involved in DNA cleavage during transformation ( 35 , 36 ), the sporulation-specific secreted nuclease NucB ( 37 , 38 ) and the minor biofilm-associated YhcR ( 39 ). When SYTOX Green culture supernatant signals were measured for strains lacking the NucA and NucB extracellular nucleases, no significant differences in extracellular DNA levels were measured compared to the wild-type culture (Supplementary Figure 3). Thus, the presence of the two major B. subtilis extracellular nucleases had no significant, measurable effect on the observed level of DNA in the culture supernatant. YhcR was not tested as it is only induced in response to the need to recover eDNA from biofilm pellicles ( 39 ). Moreover, when purified B. subtilis genomic DNA was artificially fragmented through sonication, it was found to have little effect on the SYTOX Green fluorescence levels measured (Supplementary Figure 4). Together, these results confirm that the SYTOX Green signal in the culture supernatant provides a sensitive assay for cell lysis that is largely insensitive to DNA fragmentation. Finally, to verify that the observed extracellular DNA is indeed autolysis-derived, early stationary growth phase culture samples from a strain deficient for several autolytic enzymes ( B. subtilis Δ lytABCDEF ), and from its immediate parent strain ( B. subtilis 168) were collected and analysed ( Figure 3a ). Very little DNA was observed in the culture supernatant of the autolytic mutant based on SYTOX Green fluorescence ( Figure 3b ), and no DNA was detectable in the corresponding agarose gel ( Figure 3c ). These results confirm that the DNA present in the B. subtilis culture supernatant is indeed derived from cells undergoing autolysis in a culture that, at the population average level, exhibits healthy and robust growth. Download figure Open in new tab Figure 3: Extracellular DNA present in B. subtilis culture supernatants is lysis-derived (a) Growth kinetics of B. subtilis wild type (168) and multiple autolytic enzyme -deficient derivative (Δ lytABCDEF ) strains in mSMM at 37°C. The black arrow indicates the time point of sample collection. (b) SYTOX Green fluorescence signals of corresponding culture supernatant samples. The graph depicts the mean and standard deviation from three independent biological replicates. **** = p ≤ 0.0001. (c) Agarose gel depicting DNA stained with Nancy-520 of one of the supernatant samples indicated in panel A and measured in panel b The 1 kb DNA Ladder from NEB® was used as a molecular size marker. Strains used: B. subtilis 168 (wild type) and B. subtilis KS19 (Δ lytABCDEF ). Quantification of lysis in a growing B. subtilis culture Due to the insensitivity of SYTOX Green fluorescence towards the degradation of lysis-derived DNA and the lack of significant degradation by the two major extracellular nucleases, the accumulation of extracellular DNA should serve as a cumulative measure of the lysis occurring in a culture. To estimate the extent of cell lysis required to release the observed quantities of extracellular DNA, the SYTOX Green fluorescence signal was calibrated for cell suspensions of known optical density that were lysed by sonication to release their cytoplasmic DNA into the supernatant. The respective samples were then stained with SYTOX Green, quantified using a fluorescence plate reader, and the SYTOX Green fluorescence intensities were plotted against the sample OD 600 values ( Figure 4a ). A linear correlation between the measured SYTOX Green fluorescence and corresponding OD 600 values (0-1.2) was observed ( Figure 4a, b ). This calibration facilitates a simple conversion between the measured supernatant SYTOX Green fluorescence intensity and the cumulative fraction of the cell population that has released the observed extracellular DNA through lysis. Download figure Open in new tab Figure 4: Calibration of the SYTOX Green autolysis assay (a) SYTOX Green fluorescence signals of a B. subtilis culture grown to various optical densities in mSMM and subsequently lysed by sonication to release DNA, plotted against the OD 600 of the samples prior to sonication. (b) Agarose gel showing the supernatant DNA stained with Nancy-520 in the same samples shown in panel A. The 1 kb DNA Ladder from NEB® was used as a molecular size marker. Strain used: B. subtilis prototroph (MDS23). Establishing the DiSC 3 ( 5 ) assay for measuring single-cell energy levels from B. subtilis cultures of varying growth phases and cell densities The membrane potential-sensitive carbocyanine dye 3,3′-dipropylthiadicarbocyanine iodide, DiSC 3 ( 5 ), is a well-established reporter for measuring bacterial membrane potential and, therefore, a sensitive single-cell reporter for the general bioenergetic status of various bacterial species, including B. subtilis ( 23 , 25 , 40 ). However, its use has been primarily limited to logarithmic phase, low-density cultures, and to comparing cultures with identical optical densities. The membrane potential sensitive staining by DiSC 3 ( 5 ) is due to its hydrophobic character, which allows it to diffuse across cell membranes, and the cationic charge that enables transmembrane electric fields to bias its diffusion. As a result, DiSC 3 ( 5 ) acts as a so-called Nernstian dye, establishing a gradient across the bacterial cytoplasmic membrane that scales with the membrane potential levels ( 24 , 41 , 42 ). Due to this mechanism of membrane potential sensing, the single-cell DiSC 3 ( 5 ) fluorescence levels are influenced by cell sensitivity and experimental factors that can rapidly alter the cellular metabolic state, such as changes in nutrient supply. A common experimental pitfall in this context is the washing and resuspension of cells in a nutrient-free buffer, which leads to a rapid loss of membrane potential ( 40 ). To establish a robust assay for measuring single-cell membrane potential levels for varying growth phases and cell densities, the contribution of cell density was first analysed. Accordingly, B. subtilis was grown to an OD 600 of 0.6 (exponential phase) and then diluted to an OD 600 of 0.45, 0.3 or 0.15. To prevent the re-energisation of cells through resuspension in fresh medium, cells were instead diluted in culture supernatant obtained by centrifugation of a parallel sample from the same culture. These results demonstrate that the obtained single-cell DiSC 3 ( 5 ) levels are highly sensitive towards sample optical density, with diluted samples exhibiting stronger and more homogeneous staining levels despite originating from the same culture ( Figure 5a, b ). Thus, to reliably measure and compare membrane potential in B. subtilis cultures of varying cell densities, it is necessary to equalise the sample optical density. In our experimental conditions, dilution to an OD 600 of 0.3 is sufficient to ensure homogeneous DiSC 3 ( 5 ) staining levels, but this is likely to vary if this approach is applied for different bacterial species and media. Using cell-free “spent” media obtained from corresponding cultures is strongly recommended to minimise changes in nutrient availability. Download figure Open in new tab Figure 5: Cell densities and imaging speed influence B. subtilis DiSC 3 ( 5 ) fluorescence signals (a) Phase contrast and fluorescence microscopy images of DiSC 3 ( 5 )-stained B. subtilis in mSMM in undiluted (OD 600 of 0.6) and diluted cultures (OD 600 of 0.45 to 0.15). (b) Quantification of DiSC 3 ( 5 ) fluorescence intensities for individual cells from the imaging dataset shown in panel (a) (n=100). (c) Quantification of DiSC 3 ( 5 ) fluorescence intensities for individual cells (n=90), 2-6 min after immobilisation of cells on microscope slides (application of the coverslip). Mean fluorescence intensities and standard deviation are indicated with red lines, together with P values of a one-way, unpaired ANOVA. ** = p ≤ 0.01; ns = a non-significant difference. Scale bar: 3 µm. Strain used: B. subtilis prototroph (MDS23). It has previously been shown that B. subtilis cells are affected by oxygen limitation under microscope cover slips and, as a result, gradually lose membrane potential when immobilised on a microscopy slide ( 9 , 10 ). To establish a timeframe in which DiSC 3 ( 5 ) microscopy can be applied for assessing single-cell energy levels, a B. subtilis culture was grown to an OD 600 of 0.3, stained with DiSC 3 ( 5 ), placed on the agar-coated microscope slide, covered with a cover slip, and imaged every minute up to 6 min ( Figure 5c ). At this sample cell density, B. subtilis can maintain stable membrane potential levels under the cover slip for approximately 5 min. After this time window, the membrane potential starts to decline and become more heterogeneous. In conclusion, it is possible to measure membrane potential levels reliably for relatively dilute cell suspensions (OD 600 = 0.3), provided the data is acquired within a 5-minute timeframe of immobilisation. B. subtilis wild type undergoes autolysis and exhibits strong energy level heterogeneity in glucose-based minimal medium After establishing the DiSC 3 ( 5 ) assay for measuring single-cell energy levels from B. subtilis cultures of varying growth phases and cell densities, and the SYTOX Green assay for quantifying cumulative lysis by measuring extracellular DNA, we investigated individual cell membrane potential levels and cell lysis throughout the growth cycle of B. subtilis . As shown previously ( Figure 1a ), the de-energisation of B. subtilis by collapsing the proton motive force with CCCP induces cell lysis. While the exact molecular mechanism that links cell de-energisation to cell lysis is poorly understood, it is well established that the process involves misregulation of the cell’s own autolytic enzymes ( 9 , 43 ) and is induced by numerous compounds that compromise B. subtilis membrane integrity or hamper the cell’s ability to maintain proton motive force by respiration ( 43 ). To gain greater insight into the extensive accumulation of lysis-derived extracellular DNA in an apparently healthy, well-growing B. subtilis culture, we monitored and correlated the extent of single-cell membrane potential heterogeneity and cell lysis throughout the growth cycle of B. subtilis . When B. subtilis was grown in a glucose-based minimal medium (mSMM), relatively low levels of cell lysis (cumulatively ∼5% of the cell population) were observed during the logarithmic growth phase ( Figure 6a , T 0 ). In contrast, upon slow transition from exponential to the stationary growth phase (so-called transition phase), the rate of cell lysis accelerated with ∼19% of the population having lysed by T +4 despite the continuous increase in culture optical density. This high level of background lysis was further confirmed by agarose gel electrophoresis whereby increasing amounts of extracellular DNA were detected over time ( Figure 6b ). When the corresponding single-cell energisation levels were monitored using membrane potential as a proxy, B. subtilis was found to be well energised during the exponential growth phase with little single-cell heterogeneity observed ( Figure 6b , T -4 to T -1 ). However, once the growth rate started to decline and the culture transitioned to a stationary growth phase, the average single-cell membrane potential levels gradually declined ( Figure 6b , T 0 to T +4 ). Strikingly, the cell population now began to exhibit high energy level heterogeneity with the majority of cells maintaining reasonably well-energised membranes while a sub-population of cells (∼3-6%) demonstrating near or complete de-energisation. The largest number of de-energised cells were present at T +4 h, with ∼31% of cells exhibiting very low membrane potential. Hence, there is a striking correlation between the emergence of a de-energised subpopulation and the observed increase in autolysis. It is tempting to speculate that the observed lysis is indeed due to the emergence of a de-energised cell population that ultimately undergoes autolysis. More detailed studies are needed, however, to confirm this link and decipher the physiological processes responsible for the high level of membrane potential heterogeneity observed in stationary B. subtilis cultures. Download figure Open in new tab Figure 6: B. subtilis exhibits growth phase-dependent autolysis and energy level heterogeneity (a) Growth kinetics of B. subtilis in mSMM along with the percentage of cells cumulatively lysed at specified timepoints, calculated using the calibration curve shown in Fig. 4 . (b) Agarose gel depicting DNA stained with Nancy-520 in the same lysis-derived DNA samples shown in panel A. The 1 kb DNA Ladder from NEB® was used as a molecular size marker. (c) Quantification of DiSC 3 ( 5 ) fluorescence intensities for individual cells (n=103-128) at the specific time points before and after transition (T 0 ) from exponential towards stationary phase growth. Mean fluorescence intensities and standard deviations are indicated with red lines. Strain used: B. subtilis prototroph (MDS23). DISCUSSION In this manuscript, we provide experimental guidance and protocols to extend the usability of two physiological fluorescence reporters, the voltage-sensitive dye DiSC 3 ( 5 ) and the membrane permeability indicator SYTOX Green, in the Gram-positive model bacterium B. subtilis . With the approaches detailed here, DiSC 3 ( 5 ) can be used to monitor bacterial single-cell energisation levels robustly across varying growth phases and cell densities, thereby significantly expanding its usefulness for bacterial physiological and bioenergetic studies. Furthermore, we have established how the membrane permeability indicator SYTOX Green can be used as a reporter for autolysis at the single-cell level and to quantitatively monitor cell lysis affecting a subpopulation of cells in an otherwise growing culture. While the assays are established for B. subtilis , both DiSC 3 ( 5 ) and SYTOX Green function in a variety of bacterial species ( 20 – 23 , 40 ). As a result, establishing a similar assay for other bacterial species should be relatively straightforward, building on the protocols provided here for B. subtilis . Once we had established the detailed assays for B. subtilis cultures, we encountered a surprisingly high level of membrane potential heterogeneity during late exponential and early stationary phase growth. To our knowledge, this degree of heterogeneity in energisation levels has not been previously observed. One cellular consequence of this population heterogeneity could indeed be the unexpected levels of cell lysis that affect dense B. subtilis cultures. Under these conditions, B. subtilis undergoes differentiation into various subpopulations with distinct physiological characteristics and roles ( 44 ). It is likely that the observed heterogeneity in energisation and cell lysis is connected to the developmental programmes that underpin B. subtilis differentiation, providing sources of nutrients under conditions where exogenously supplied nutrients are exhausted. While we are currently actively studying the underlying molecular mechanism and its consequences for B. subtilis physiology, details regarding this underappreciated biological phenomenon are beyond the scope of this methods-focused manuscript. DATA AVAILABILITY Source data for all figures and graphs presented in the manuscript will be made available via Newcastle University’s research data repository via https://doi.org/10.25405/data.ncl.29924237 . The strains and plasmids are available upon request to H.S. AUTHOR CONTRIBUTIONS M.D.S and M.H. performed the experiments, M.D.S, T.v.R, C.R.H and H.S designed experiments and helped with data interpretation, T.v.R, C.R.H and H.S conceived and supervised the project, M.D.S, C.R.H and H.S wrote the manuscript. ACKNOWLEDGMENTS This research was supported by the UK Biotechnology and Biological Sciences Research Council grants BB/S00257X/1 and BB/M011186/1, and the Barbour Foundation. We thank James Grimshaw for his support with image analysis. For the purpose of open access, the authors have applied a Creative Commons Attribution (CC BY) licence to any author-accepted manuscript version arising from this submission. Funder Information Declared Biotechnology and Biological Sciences Research Council , BB/S00257X/1 , BB/M011186/1 Barbour Foundation Footnotes https://doi.org/10.25405/data.ncl.29924237 REFERENCES 1. ↵ Arnaouteli S , Bamford NC , Stanley-Wall NR , Kovács ÁT. 2021 . Bacillus subtilis biofilm formation and social interactions . 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