The neutron scattering signal from a cell reflects the sum of contributions from all cellular components, here, taken to be water, protein, RNA, DNA, carbohydrate, and lipid (Fig 1a). Each of these has a characteristic ρ (Fig 1b), and from their relative abundances in B. subtilis,[33] we estimated the total scattering from the organism as a function of D₂O concentration in the aqueous environment. Water accounts for approximately 85% of the cellular volume and exchanges rapidly across the membrane. Therefore, immersion of cells in deuterated buffer changes the neutron contrast, ergo the net scattering, with the relative contributions of the different molecular species varying as a function of D₂O concentration (Fig 1c; S2 Fig).

We then measured the total scattering from B. subtilis cultured in standard M9 minimal medium (H-M9) and resuspended in phosphate buffered saline (PBS) at different D₂O concentrations. The observed scattering corresponded closely to predictions of the simple compositional model, as shown in Fig 1c. Note that the cells scatter strongly over the entire range of percent D₂O, and the total signal at any concentration of D₂O reflects contributions from the multiple classes of biomolecules present in the cell. Structural analysis based on the overlapping signals is impractical, if not impossible.

To isolate a meaningful scattering signal from the membrane, it was first necessary to suppress neutron contrast from the entire cell by manipulating its H/D composition. This objective was achieved by culturing B. subtilis in deuterium-enriched M9 minimal medium (D-M9, prepared in 90% D₂O with H-glucose as the carbon source). Due to metabolic H/D exchange, deuterium from the growth medium becomes permanently incorporated into the carbon skeletons of biosynthetic molecules. Overall, skeletal deuteration was approximately 70% (S4 Fig, Tables B and C in S1 Text), which was predicted to create a near-contrast matched condition for all biomolecules when the cells were immersed in approximately 85% D₂O buffer (Fig 2a). This expectation was borne out in the scattering experiment, where a strong reduction in the total scattering was observed at approximately 85% D₂O (Fig 2b, open circles; S2 Fig).

With contrast suppressed, the next step was to reintroduce contrast specifically into the membrane by providing exogenous FAs for incorporation into the membrane phospholipids. Membrane FAs, after conversion to FA methyl esters (FAMEs), are readily analyzed for structure and isotope substitution by gas chromatography/mass spectrometry (GC/MS). As shown in Fig 2c (top), B. subtilis uses a mixture of 7 main FAs, all of which are saturated (have no double bonds), and all but one of which are branched.[34,35] Initial supplementation experiments with wild-type B. subtilis showed that exogenous FAs were not incorporated into the membrane lipids intact, so the native pathways for both catabolism and anabolism of FAs had to be blocked (Fig 2d).

Catabolism was blocked genetically by the deletion of yusL, the gene encoding a critical enzyme in β-oxidation, enoyl-CoA hydratase.[36] Anabolism was blocked chemically using cerulenin, an irreversible inhibitor of β-ketoacyl-ACP synthase that suppresses de novo FA biosynthesis.[37] This combination resulted in conditional FA-dependent growth (demonstrated in S5 Fig). Cerulenin-induced growth inhibition can be rescued with a mixture of just 2 of the native complements of FAs—palmitic acid (normal-hexadecanoic acid [n16:0]) and 12-methyltetradecanoic acid (anteiso-pentadecanoic acid [a15:0]).[38] These 2 constitute a minimal set of 1 high-melting (n16:0) and 1 low-melting (a15:0) FA, with which the cell can regulate the fluidity and structure of its membrane. Unlabeled (H) and perdeuterated (D) forms of these 2 FAs were then used in various mixtures to tune neutron contrast in the membrane.

Growth of B. subtilis ΔyusL in D-M9 medium did not alter the FA composition of the membrane (Fig 2c, upper and middle panels; see S4 Fig for peak assignments and Table B in S1 Text for peak areas and deuteration analysis). However, when cerulenin-treated ΔyusL cells were grown in the same D-M9 medium, supplemented with H-n16:0 and H-a15:0, their membrane FAs were found to consist exclusively of these 2 FAs, with no deuterium incorporated from the medium (Fig 2c, bottom panel). The total neutron scattering from these cells showed a large increase in scattering at 85% D₂O (marked by the blue arrow in Fig 2b), which can be attributed entirely to the H-FAs incorporated into the membrane phospholipids.

Having imparted contrast to the membrane (Fig 3a), we used SANS to determine the transverse membrane structure by recording the scattered intensity, I(q), as a function of the scattering wavevector, q. Cells for this experiment were cultured as described above in D-M9 medium, supplemented with cerulenin and H-n16:0 and H-a15:0 to label the membrane, then transferred to 85% D₂O buffer. Because these are more time-consuming experiments, the cells were resuspended in an 85% D₂O buffer supplemented with glucose, Mg²⁺, and cerulenin. These additives prevent autolysis and preserve the membrane potential,[39,40] such that cells in suspension remained >90% viable over a period of 4 h at 25°C, as determined by direct cell counts, optical density measurements, and live/dead staining (S6 and S7 Figs). The residual background was recorded using cerulenin-treated ΔyusL cells, which were fed a mixture of FAs contrast-matched to 85% D₂O (a15:0 and n16:0, each 30% H and 70% D), and which do not contribute substantially to the net neutron scattering signal.

In SANS, the shape of the I(q) versus q data describes the structure of the sample on the order of Ångstroms to approximately 100 nm. Formally this data is modeled using a lamellar form factor representing the acyl core of the bilayer, with the 2 identical head group regions on either side, contrast-matched to the scattering density of the surrounding cellular environment. The lamellar form factor is characterized by a q⁻² dependence at low-q, arising from the 2D membrane surface of the entire bacterium, transitioning to a q⁻⁴ dependence typical of 3D objects with smooth surfaces and sharp interfaces—more detail is available in the literature[41–44] and the materials and methods.

Subtraction of the background from the sample scattering revealed a pure membrane spectrum, which displayed a lamellar form factor characteristic of a lipid bilayer (Fig 3b), with the expected q⁻² dependence at low-q (dashed line). A fit of the data (solid red line) revealed the average membrane hydrophobic thickness (2D_C) to be 24.3 ± 0.9 Å at 25°C (S8 Fig). The hydrophobic thickness of the living B. subtilis membrane is thus comparable to that of synthetic phosphatidylcholine (PC) bilayers, such as dimyristoyl PC (2D_C = 25.7 Å at 30°C) or 1-palmitoyl-2-oleoyl PC (2D_C = 28.8 Å at 30°C).[46] It is also comparable to that of purified basolateral plasma membranes from rat hepatocytes (approximately 26 Å), as measured by small-angle X-ray scattering,[6] highlighting the conservation of membrane structure across animal and eubacterial kingdoms.

Finally, we sought to examine the lateral structure within the membrane to determine whether the lipids in the B. subtilis membrane are uniformly mixed or display nanoscopic organization, as predicted by the lipid raft hypothesis.[21] Canonical mammalian lipid domains are believed to be enriched in the high-melting lipids, cholesterol and sphingomyelin. Bacteria generally lack these lipids, but are nonetheless believed to have lipid domains[47–53] formed from lipid species playing analogous roles. [54,55] The expected hallmark of lipid domains, in any system, is the lateral separation of higher- and lower-melting lipids with associated proteins into distinct phases.

We recently introduced a contrast-variation strategy to detect lateral lipid organization in synthetic vesicles using SANS[28,29]. This strategy relies on differential H/D labeling of the lipid phases to control contrast in the plane of the membrane. With a suitable labeling strategy, the separated phases can be either contrasted or matched with respect to each other and the buffer. A particularly important case is where a mixture of lipids has an average contrast matching that of the buffer, but partitions into H- and D-enriched phases that contrast each other and the buffer. Experimentally, uniform mixing creates a null-scattering condition, whereas phase separation in the plane of the membrane (domain formation) induces neutron contrast and an observable signal in the SANS spectrum. In S9 Fig and Table F in S1 Text we present a series of similar SANS experiments which show how neutrons are sensitive to contrast in the plane of the bilayer and how neutron contrast can be controlled by thermally induced mixing of the 2 phases or by selecting specific isotopic mixtures which result in contrast-matched phases.

The in vivo adaptation of this strategy compares scattering from cells with 2 different H- and D-FA mixtures that are, on average, contrast-matched to the medium (Fig 4a). In the control mixture, there can be no contrast regardless of whether or not the membrane lipids are uniformly mixed, because all species are present at the same H/D ratio. However, in the experimental mixture, de-mixing among lipids creates contrast as described above, producing a measurable increase in the scattered intensity as a result of local inhomogeneities in the H/D distribution.

We implemented this strategy in cerulenin-treated B. subtilis ΔyusL cells using an experimental mixture of 100% D n16:0 (high-melting) and 40/60 H/D a15:0 (low-melting). When corrected for the relative abundances of these FAs in the cell (S10 Fig), this mixture creates an average membrane contrast matching the 30/70 H/D FA ratio (a15:0 and n16:0, each 30% H and 70% D) used for the control mixture (inset to Fig 4b) as well as the rest of the cellular components, including the 85% D₂O buffer. SANS spectra from B. subtilis fed the experimental mixture reproducibly displayed an excess scattering over the q range 0.015–0.2 Å⁻¹ compared to cells fed the control mixture (results from a repeat experiment are shown in S11 Fig). As the only source of contrast in the experiment was the membrane H/D FA pool, this result supports the notion that there is lateral de-mixing of lipids containing the high- and low-melting FAs on a length scale ℓ of 3–40 nm (ℓ = 2π/q) (Fig 4c), consistent with the lipid raft hypothesis.

Deuterium oxide (99.9% D) and algal amino acids (unlabeled and uniformly D-labeled, with an isotopic purity of 98%) were obtained from Cambridge Isotope Laboratories. Palmitic acid (H-n16:0) and palmitic acid-d₃₁ (D-n16:0) were obtained from Sigma–Aldrich. 12-Methyltetradecanoic acid (H-a15:0) was purchased from Sigma–Aldrich or prepared from s-butyl magnesium chloride (Sigma–Aldrich) and 11-bromoundecanoic acid (Sigma–Aldrich) according to the method of Baer and Carney[64] and purified by vacuum distillation. Perdeuterated D-a15:0 was prepared from H-a15:0 through 3 cycles of H/D exchange with D₂O catalyzed by 10% Pt/C at 220°C as described by Yepuri et al.,[65] followed by chromatography on silica gel and vacuum distillation. The final product was chemically homogeneous and had an isotopic purity of 99%, as determined by analysis of its derived methyl ester by GC/MS. Cerulenin was obtained from Alfa Aesar (Tewskbury, MA) and stored in the dark at –80°C as a solid. FA-free bovine serum albumin (BSA, catalog number A8806) was obtained from Sigma–Aldrich (St. Louis, MO). All other materials were obtained from commercial suppliers and used as received.

B. subtilis 168 (parent strain) and a ΔyusL mutant (strain BKE32840) were obtained from the Bacillus Genetic Stock Center (The Ohio State University, Columbus, OH). The ΔyusL strain lacks enoyl-CoA hydratase/3-hydroxyacyl-CoA dehydrogenase activity and is severely deficient in its ability to degrade exogenous long-chain FAs;[36] therefore, this strain was used for all experiments where exogenous FAs were supplied in the cultivation medium. General growth and maintenance of B. subtilis was performed in either Luria–Bertani (LB) rich medium or M9 minimal medium with 2% glucose supplemented with 5 mM l-tryptophan. Solid media were prepared by the addition of 1.5% Bacto or Noble Agar (Difco) to LB or M9 medium, respectively. Erythromycin was added to 0.5 μg/mL for routine maintenance of BKE32840. Cultures were incubated at 37°C, and liquid cultures were aerated by shaking at 250 rpm. Where applicable, supplemental FAs were added to a final concentration of 8 mg/L each of a15:0 and n16:0 from 25 mg/mL stock solutions in ethanol, along with 10 g/L of FA-free BSA as a carrier to aid solubility. Cerulenin (10 mg/mL in ethanol stock solution) was prepared fresh and added immediately prior to inoculation. The required final concentration was determined empirically and varied by supplier and batch. For the work described here, cerulenin from Alfa Aesar was used at a final concentration of 50 μg/mL, which was sufficient to fully suppress endogenous FA synthesis, as judged by inhibition of growth, and rescued by exogenous FA (S5 Fig).

Partially deuterated cells were grown in M9, prepared using 90% v/v D₂O and H-glucose. This medium produced approximately 60%–70% deuteration of the carbon skeletons in biosynthetic molecules (analysis described below). B. subtilis 168 and BKE32840 were adapted to growth in M9-Gluc + 5 mM l-tryptophan prepared with 90% (v/v) D₂O by serial passage (1:100 inoculum) in media prepared with increasing concentrations of D₂O (H₂O:D₂O 100:0, 50:50, and 10:90). Cerulenin-treated, FA-supplemented cells were grown starting from a culture of untreated, unsupplemented cells. The starter culture (OD₆₀₀ 0.8–1.0) was diluted 1:20 in cerulenin/FA medium to an OD₆₀₀ of ca. 0.05, incubated for growth to an OD₆₀₀ of 0.8–1.0, then passaged by dilution (1:20) into fresh medium. FA composition was monitored at each passage by GC/MS; 5–8 passages were required to clear the native FAs and adapt the cells to the FA-dependent condition. Control cultures without supplied FA (no-growth controls) were monitored in parallel for each experiment to ensure that the cerulenin remained active during the incubation period. Optical densities were determined at 600 nm using a Synergy Mx plate reader (BioTek, Winooski, VT, USA), using a 96-well microtiter plate and 300-μL well volumes.

For contrast experiments (Figs 1 and 2, and S2 Fig), B. subtilis 168 cells were resuspended in PBS (10 mM Na₂HPO₄, 1.8 mM KH₂PO₄, 137 mM NaCl, and 2.7 mM KCl), prepared either with H₂O or D₂O (H-PBS or D-PBS, respectively), then mixed in appropriate proportions. For all H cells (Fig 1c), 250 mL of fresh culture (OD₆₀₀ = 1.5) in H-M9 medium was split into 2 parts, which were processed in parallel. Cells were harvested by centrifugation at 6000 × g for 20 min at 4°C, then washed by centrifugation/resuspension with 3 × 10 mL of H- or D-PBS, allowing 5 min for equilibration at each step. The washed cell pellets were then resuspended in 11 mL of the same buffer to provide a cell concentration approximately 50 mg/mL wet weight, equivalent to approximately 10 mg/mL dry weight. For deuterated cells (Fig 2), the same procedure was followed except that a 125-mL culture grown in D-M9 prepared with 90% D₂O was used, providing a final cell concentration of approximately 5 mg/mL dry weight. Cell suspensions were loaded into quartz “banjo” cells (diameter 22 mm, path length 1 mm) for study by SANS. Measurements were made at 37°C, using a single detector position, and the total scattering was analyzed as described below.

For membrane structural analyses (Figs 3 and 4, S7 and S11 Figs), FA-fed B. subtilis 168 ΔyusL cells were grown in M9-Gluc prepared with 90% D₂O. Cells from a 35-mL culture were harvested at an OD₆₀₀ of 0.5, washed with 3 × 3 mL of PBS, prepared with 85% D₂O, and resuspended in 1 mL of the same buffer, providing a final cell concentration of approximately 5 mg/mL dry weight. Because of the long data collection times required (up to 4 h), glucose (0.1% w/v), MgSO₄ (10 mM), and cerulenin (50 μg/mL) were added to the final resuspension buffer, the pH was reduced to 6.8, and the measurements were made at 25°C to prolong cell viability and minimize autolysis.[39,40] As discussed below under cellular viability and integrity and shown in S6 and S7 Figs, cells remained >90% viable and displayed consistent SANS spectra over a period of 4 h under these conditions.

Lipids were extracted from cells and characterized for FA content by GC/MS of derived FAMEs (schematic description provided in S4 Fig). Total lipid extracts were prepared using a modification[66] of the method of Bligh and Dyer.[67] In brief, cells were pelleted by centrifugation at 6000 × g for 15 min, followed by 3 washes in 1% (w/v) NaCl. Samples were lyophilized in 10 mL glass test tubes with Teflon-faced screw caps, to each of which was sequentially added 0.5 mL of chloroform, 1 mL of methanol, and 0.4 mL of water, with vigorous agitation at each stage. This mixture forms a single phase and was left to stand for 18 h at room temperature with occasional agitation. After 18 h, phase separation was induced by the addition of 0.5 mL of chloroform and 0.5 mL water. The lipids were recovered from the lower chloroform phase by evaporation of the solvent in a new 10-mL glass test tube under an argon stream. FAMEs were prepared by acidic methanolysis of dried lipid extracts or intact cells.[68] Solvents, if present, were evaporated under a stream of argon prior to the addition of 1 mL of concentrated HCl/methanol (10% v/v). The test tube was then purged with argon, sealed, and heated to 85°C for 2 h. After cooling, 1 mL of water and 1 mL of hexane were added, and the contents were vortex-mixed. After phase separation, a portion (approximately 700 μL) of the upper phase was taken out for GC/MS analysis.

Cellular amino acids were analyzed by GC/MS, as described by Dauner and Sauer.[69] Cells from 10 mL of culture at OD₆₀₀ ≈ 0.7 were harvested by centrifugation, washed 3 times with water by resuspension/centrifugation, and stored frozen. For analysis, cells were resuspended in 1 mL of water in a microcentrifuge tube and lysed by sonication. After centrifugation at 14,000 × g for 15 min, a 500-μL portion of the supernatant was transferred to a glass vial and mixed with 1.5 mL of 6 M HCl. Standard solutions were prepared from H- or D-algal amino acid mixtures and processed in the same manner. The vials were sealed with Teflon-lined caps and heated at 110°C for 24 h to hydrolyze protein, after which the volatiles were removed by rotary evaporation. The hydrolysates were resuspended in tetrahydrofuran (100 μL) and N-tert-butyldimethylsilyl-N-methyltrfluoroacetamide (100 μL) and then heated to 60°C for 1 h to produce tert-butyldimethylsilyl amino acid derivatives suitable for GC/MS analysis. Samples were diluted with hexane 1:10 (v/v) prior to analysis.

GC/MS analysis was performed using an Agilent 5890A gas chromatograph with a 5975C mass-sensitive detector operating in electron-impact mode (Agilent Technologies, Santa Clara, CA). The instrument was equipped with an HP-5ms capillary column (30 m long, 0.25 mm outside diameter, and 0.25 μm coating thickness) using helium at 1 mL/min as the carrier gas. Samples of 1 μL were introduced using splitless injection at an inlet temperature of 270°C. FAMEs were eluted using a temperature program of 2 min at 60°C; 20°C/min to 170°C; 5°C/min to 240°C; and 30°C/min to 300°C for 2 min. Derivatized amino acids were eluted with a temperature program of 2 min at 100°C; 10°C/min to 280°C for 2 min.; and 25°C/min to 325°C for 2 min. Peak assignment, integration, and mass spectral analysis were performed using the instrument's ChemStation Enhanced Data Analysis software and the NIST mass spectral database. Peaks for deuterated compounds were identified on the basis of retention times and spectral comparison with nondeuterated compounds. The extent of deuteration was assessed by determining the gain in molecular mass for parent ions of FAMEs (Table B in S1 Text) or of selected fragment ions for amino acids (Table C in S1 Text).

SANS data were collected on Bio-SANS[70] and the Extended Q-range Small Angle Neutron Spectrometer (EQ-SANS)[71] at the at the High Flux Isotope Reactor and the Spallation Neutron Source, respectively, both located at Oak Ridge National Laboratory. Two-dimensional scattering data from both instruments were reduced using the Mantid[72] software and normalized to a porous silica standard to establish an absolute scale, and corrected for pixel sensitivity, dark current, and sample transmission. Background scattering was subtracted from the 1D intensity versus q, which is defined as: q= 4πsin(θ)/λ (1) where λ is the neutron wavelength and 2θ is the scattering angle relative to the incident beam. At EQ-SANS, the data were collected in 60 Hz mode with 2 instrumental configurations: 1.3-m sample-to-detector distance with 4–7 Å neutrons (q = 0.050–0.4 Å⁻¹) and 4.0 m sample-to-detector distance with 10–13.4 Å neutrons (q = 0.009–0.07 Å⁻¹), yielding a total q-range from approximately 0.009 to 0.4 Å⁻¹. At BioSANS, 6-Å neutrons were used at 2 sample-to-detector distances, 2.5 m (q = 0.050–0.30 Å⁻¹) and 15.3 m (q = 0.005–0.060 Å⁻¹), yielding a total q-range from 0.005 to 0.30 Å⁻¹. Collection times did not exceed 4 h, at which point cells were determined to be better than 90% viable, as shown below in cell viability and sample integrity (S6 and S7 Figs). The scattering data from the cells demonstrate no statistically significant change in scattered intensity after 4 h (S7 Fig).

Model lipid mixtures (S9 Fig) were produced using synthetic lipids from Avanti Polar Lipids (Alabaster, AL) and prepared as follows: lipids were dissolved in chloroform, dispersed as a film by evaporation in a 20-mL scintillation vial, and dried overnight under vacuum (>6 h). The lipid films were then rehydrated in the isotopically appropriate solvent (H₂O, D₂O, or a combination thereof) and subjected to 5 freeze-thaw cycles prior to extrusion through 100-nm pore-diameter track-etched polycarbonate membrane filters. The final concentration of vesicles for scattering measurements was 10 mg/mL.

Data used for the contrast experiments (Figs 1 and 2) were collected on EQ-SANS for 0.009 Å⁻¹ < q < 0.06 Å⁻¹. The data were evaluated as the Porod invariant: Q*=∫q2I(q)dq=2πI(0)Vp (2)

The proportionality of Q* to the total scattering intensity I(0) makes it a useful metric for comparison to structure-independent estimates of I(0) based only on composition and scattering length density, ρ. V_p is the Porod volume. To analyze the acyl thickness of the bilayer, the data were modeled using the expression:[73] I(q)=NVVs2(ρm−ρs)2〈|F(q)|2〉 (3) for an arbitrary number of bilayers, N, of volume, V_s, with scattering length density ρ _s, in a solvent of ρ_m, where F(q) is the form factor describing the lamellar shape, and (ρ_m − ρ_s), is the contrast term. The scattering law used to model the data was the lamellar form factor representing the hydrocarbon core of the bilayer, with 2 identical head group regions on either side, with scattering densities matching that of the surrounding cellular environment. Fitting of the SANS data was performed in SASview.[74]

Estimates of total scattering from B. subtilis (Figs 1 and 2) were made as follows. A neutron scattering signal arises when there is a difference in neutron scattering length density, ρ, between a species, s, and the medium in which it resides in, m. The difference, (ρ_m−ρ_s), is called contrast, and the scattered intensity is proportional to its square. For any species, ρ=(Σinbi)/V (4) where n is the number of atoms, b is the coherent neutron scattering length for each atom, and V is the volume of the species. Because each class of biomolecule (i.e., protein, lipid, carbohydrate, RNA, or DNA) has a nearly constant chemical composition and density, ρ is well approximated as a single value for each class (Table A in S1 Text).[30]

Deuterium labeling increases the ρ of an unlabeled molecule (ρ_H) because the scattering length b is greater for D than H (6.67 versus –3.74 fm).[31] In biomolecules, the hydrogen atoms can be assigned to 2 categories, those bound to carbon (CH), which do not exchange with water, and those bound to heteroatoms (XH, X = N, O, or S), which do exchange with water. The fractions in the 2 categories are consistent within each class of biomolecule, which allows for the definition of the terms, Δρ_CD and Δρ_XD, that represent the increase in ρ resulting from the complete deuteration of each category. For a deuterated biomolecule: ρ=ρH+fXDΔρXD+fCDΔρCD (5) where ρ_H is ρ for the unlabeled (all-H) molecule, and f_XD and f_CD are the fractions of deuterium substitution in the XH and CH categories, respectively. Estimates (Figs 1b and 2a) were generated using Eq 5 and the values for ρ_H, Δρ_XD, and Δρ_CD in Table A in S1 Text. The differing slopes for each class of biomolecule reflect differences in Δ_XD.

The total scattering from a cell, I_cell(0), is the sum of the scattering contributions from all of the cell’s biomolecular species, given by the relation Icell(0)∝Σsχs(ρm−ρs)2 (6) where χ_s is the volume fraction of each species s, ρ_s is its neutron scattering length density, and ρ_m is the average neutron scattering length density of the medium (all species s, including intracellular water). When ρ_s = ρ_m, the species is contrast-matched and thus effectively invisible to neutrons, provided the medium is uniform. The scattering attributable to an individual species j is given by: Ij(0)∝0.5χjΣsχs(ρj−ρs)2 (7)

In an undeuterated cell, the χ_j for biomolecules is sufficiently high and the ρ_j is sufficiently different that the surrounding medium is not truly uniform. As a result, the total scattering for a given species j reflects interspecies contrast and is not completely nulled at the nominal contrast matchpoint defined by the average of the medium (χ_j = χ_m). However, judicious deuteration can be used to converge ρ for water and most biomolecules (cf. Figs 1b and 2a), effectively suppressing the interspecies contributions.

Knowledge of the cell’s composition (Table A in S1 Text) allows one to estimate the total cellular scattering—broken down by biomolecular species as a function of deuteration in the solvent and in the CH skeletons of the various biomolecules (Figs 1c and 2d). Approximately 80% of the dry mass of a B. subtilis cell is made up of protein (53%), RNA (18%), DNA (2.6%), lipid (5.2%), and carbohydrate (2.8%).[33] The remaining mass—which was neglected in our estimates—consists of small organic molecules, such as amino acids, cofactors, and nucleotides plus inorganic material. These data were used with Eqs 6 and 7 to calculate the predicted total scattering in Fig 1c.

Estimated scattering for cells grown in D-M9 (Fig 2b) relied on analyses of lipids and proteins extracted from B. subtilis under relevant conditions to adjust ρ according to Eq 5. The net deuteration of the FA pool in the absence of cerulenin and supplemental FAs was 68.5%, with deuteration of individual FAs ranging from 66% to 71% (Table B in S1 Text). The deuteration of the amino acid pool was more variable and had a lower average deuteration of 60% (Table C in S1 Text), which is similar to what would be expected in Escherichia coli.[75] A value of 70% was assumed for other biomolecules, and the extent of deuteration at water-exchangeable positions was assumed to match that of the medium.

Cell viability was evaluated using optical density measurements, manual cell counts with a hemocytometer, and fluorescent live/dead staining with the BacLight Bacterial Viability Kit (Catalog number L7012, Molecular Probes, Eugene, OR). The live/dead assay uses a mixture of 2 fluorescent nucleic acid stains (SYTO 9 and propidium iodide), which stain live cells green and dead cells with compromised membranes, red. Fluorescence micrographs of stained cells were acquired with a Zeiss Axioskop 2 Plus and analyzed with ImageJ[76] to count cells using green and red fluorescence channels. Nonirradiated suspensions of the ΔyusL strain prepared identically to the samples used for neutron beam experiments were used as controls. Cell suspensions were incubated in a sealed cuvette at 25°C, and optical density (OD₆₀₀) was recorded at 1-min intervals over a period of 24 h (S6a Fig). Direct cell counts and live/dead staining of the nonirradiated samples were performed immediately after processing and at 4 h and 24 h (S6b Fig). These experiments showed close correspondence among the 3 measures of viability and that cells remained 90% viable over 4 h and 50%–60% viable over the course of 24 h under the conditions of the experiment. Irradiated cell suspensions were taken from the neutron beam after data collection and allowed to decay for approximately 30 min, at which time they were subjected to a radiological survey. Radiological safety protocols do not permit timely removal of cell suspensions for microscopic examination. Instead, optical density measurements of the cells at 4 h and 24 h were carried out using a Shimadzu UV-2700 UV–Vis spectrophotometer (S6a Fig). FA stability was monitored over the course of the 4 h it took to collect a complete SANS data set using GC/MS as described above for nonirradiated samples (S7 Fig). Finally, cell stability in the neutron beam was assessed by repeating the SANS measurement 3.5 h after the cells were put in the beam showing no change in the scattered intensity (S7 Fig).