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Applied and Industrial Microbiology
Research Article
31 January 2023

Minimizing the Lag Phase of Cupriavidus necator Growth under Autotrophic, Heterotrophic, and Mixotrophic Conditions


Cupriavidus necator has the unique metabolic capability to grow under heterotrophic, autotrophic, and mixotrophic conditions. In the current work, we examined the effect of growth conditions on the metabolic responses of C. necator. In our lab-scale experiments, autotrophic growth was rapid, with a short lag phase as the exponential growth stage was initiated in 6 to 12 h. The lag phase extended significantly (>22 h) at elevated O2 and CO2 partial pressures, while the duration of the lag phase was independent of the H2 or N2 partial pressure. Under heterotrophic conditions with acetate as the organic substrate, the lag phase length was short (<12 h), but it increased with increasing acetate concentrations. When glucose and glycerol were provided as the organic substrate, the lag phase was consistently long (>12 h) regardless of the examined substrate concentrations (up to 10.0 g/L). In the transition experiments, C. necator cells showed rapid transitions from autotrophic to heterotrophic growth in less than 12 h and vice versa. Our experimental results indicate that C. necator can rapidly grow with both autotrophic and heterotrophic substrates, while the lag time substantially increases with nonacetate organic substrates (e.g., glucose or glycerol), high acetate concentrations, and high O2 and CO2 partial pressures.
IMPORTANCE The current work investigated the inhibition of organic and gaseous substrates on the microbial adaption of Cupriavidus necator under several metabolic conditions commonly employed for commercial polyhydroxyalkanoate production. We also proposed a two-stage cultivation system to minimize the lag time required to change over between the heterotrophic, autotrophic, and mixotrophic pathways.


Biodegradable plastics are gaining increasing attention as a sustainable alternative to petroleum-based chemical plastics because of their biodegradability in the environment. Polyhydroxyalkanoates (PHA) is a biodegradable polymer produced by various microorganisms with characteristics comparable to those of petrochemical plastics (1). Among over 300 identified PHA accumulators, only a few microorganisms, such as Cupriavidus necator (previously Wautersia eutropha [2], Alcaligenes eutrophus [3, 4], and Ralstonia eutropha [58]), can be employed for commercial and large-scale PHA production (9, 10) because of their high PHA accumulation capabilities. For instance, C. necator cells can accumulate up to 90% (wt/wt) of their dry cell weight as PHA (1113).
C. necator can utilize inorganic carbon sources such as CO2 and CO, as well as organic substrates (1, 14, 15). As a hydrogen-oxidizing bacterium, C. necator can obtain the energy required for CO2 fixation by oxidizing H2 (i.e., the electron donor) and reducing O2 (i.e., the terminal electron acceptor) (14, 16). The recommended gas mixture composition to attain sufficient cell growth has a volume-based H2/O2/CO2 ratio of 7:2:1 (16). O2 concentrations above 0.30 atm are known to inhibit cell growth (17). In addition, hydrogen concentrations lower than 0.04 atm resulted in low biomass yields (18). Although the inhibiting effects of gaseous substrates on cell growth were previously reported, the inhibition of microbial adaption (i.e., lag phase) has not yet been investigated in the literature. Therefore, one of the main objectives of the current work was to examine the impact of the partial pressure of individual gases (e.g., O2, CO2, H2, and N2) on the lag phase of C. necator cells. We also focused on identifying parameters affecting the lag duration under autotrophic conditions so that we could optimize the gas-to-solution ratio and the frequency of gaseous substrate injection.
C. necator cells can also grow and accumulate PHA by utilizing organic substrates as the energy and carbon sources (4, 19, 20). C. necator was examined to grow and accumulate PHA on various organic substrates such as acetate (21, 22), glucose (23), glycerol (24, 25), valerate (26), and other waste organics (26, 27). The elevated concentrations of organic substrates may inhibit the heterotrophic growth of C. necator. For example, acetic acid concentrations higher than 0.50 g/L were previously reported to stop microbial growth (22). Additionally, the lag phase increased significantly with increasing concentrations of volatile fatty acids (9). Therefore, we investigated the impact of various concentrations (1 to 10 mg/L) of several organic substrates (e.g., acetate, glucose, glycerol, and organic mixtures) on the lag phase lengths and biomass yields of C. necator.
C. necator can also consume organic and inorganic carbon substrates simultaneously (3, 9). In previous studies, the C. necator mixotrophic growth on lactate was more rapid than individual autotrophic or heterotrophic growth, and the mixotrophic biomass yields were significantly higher than in the heterotrophic experiments for the same amount of lactate consumed (28). However, C. necator cells were not able to grow mixotrophically on some substrates, such as pyruvate (29). The mixotrophic growth of C. necator on various organic substrates (e.g., acetate, glucose, and glycerol) and their mixtures has not been systematically explored in previous studies. Thus, another novelty of this study was to evaluate the mixotrophic growth of C. necator for various combinations of multiple organic and mixotrophic conditions. In this work, we also investigated the ability of C. necator to transit between the metabolic pathways (i.e., subsequent transition from autotrophic to heterotrophic or vice versa), with a focus on the lag time (i.e., adaptation time).


Autotrophic growth.

C. necator cells grew rapidly with short lag phases (<12 h) when cultivated with the standard gas mixture (H2/O2/CO2 = 7:2:1) (Fig. 1A). The highest optical densities were achieved in 22 h, indicating fast fixation of carbon dioxide. In addition, the biomass concentrations were dependent on the provided gas volumes (Fig. 1B). As a result, the biomass concentration at 24 h was 510 ± 25 g/L chemical oxygen demand (COD) at the highest ratio between the gas (14 mL) and growth solution (3 mL) volumes. Furthermore, the total biomass was linearly correlated with the provided gas volumes (Fig. 1C), signifying the dependence of microbial growth on the gas mixture volumes.
FIG 1 (A) Autotrophic C. necator growth at different ratios between gas (14 to 5 mL) and growth solution (3 to 12 mL) volumes (all experiments were conducted in duplicate [n = 2]). (B) Maximum autotrophic net biomass concentration at different gas volumes after 24 h. (C) Relation between autotrophic biomass (concentration [mg COD/L] × solution volume [mL]) and the supplied gas volumes (3 to 12 mL). In the control vials, the autotrophic growth solution was used under the gas mixture (H2/O2/CO2 = 7:2:1) without microbial inoculation.
Higher optical densities were also achieved by increasing the frequency of gas addition to the headspace (see Fig. S1 in the supplemental material). However, additional injections of an individual gas (i.e., H2, O2, or CO2) did not enhance the biomass concentration substantially (Fig. S1), indicating the necessity of maintaining specific gas ratios to attain the highest cell productivity and avoid gas limitation (3032).

Impact of individual gas partial pressures on autotrophic growth.

Higher oxygen and carbon dioxide partial pressures severely inhibited the autotrophic growth of C. necator and resulted in longer lag phases (Fig. 2). The highest optical density at 600 nm ([OD600] = 0.78) was achieved with the standard gas mixture at a total pressure of 1 atm, indicating that the standard gas composition (H2/O2/CO2 = 7:2:1) and pressure were optimal for autotrophic growth. C. necator growth was suppressed when the value of PO2 exceeded 0.70 atm (Fig. 2A). Ishizaki and Tanaka reported that lower oxygen concentrations (<0.21 atm) were preferable to attain high C. necator growth rates, and growth almost ceased when the value of PO2 exceeded 0.30 atm (17), which matches well with the current results.
FIG 2 Impact of individual gas pressures on C. necator autotrophic growth. The initial gas mixture was H2/O2/CO2 = 7:2:1, and then the headspace pressure was increased up to 3.0 atm using O2 (A), CO2 (B), H2 (C), or N2 (D) (all experiments were conducted in duplicate [n = 2]). In the control vials, the autotrophic growth solution was used under 1.0 atm of the gas mixture (H2/O2/CO2 = 7:2:1) without microbial seed.
Similarly, a PCO2 value higher than 0.10 atm resulted in slower C. necator growth with longer lag phases (>12 h) (Fig. 2B). For example, the lag phase (37 h) at PCO2 = 2.10 atm was much longer than that at PCO2 = 0.10 atm (<12 h). In addition, the highest OD600 value at PCO2 = 2.10 atm (after 60 h) was 58% lower than that at PCO2 = 0.10 atm (after 22 h). It should be noted that the lowest pH value was 6.02 at PCO2 = 2.10 atm at the end of the experiments (Fig. S2), due to the buffering capacity of phosphate in the medium solution. The optimal pH range for the maximal C. necator growth rate under autotrophic conditions was reported to be between 6.4 and 6.9 (19), indicating that the low pH values resulting from elevated CO2 partial pressures yielded longer lag phases during autotrophic growth (19).
Applying high H2 or N2 partial pressures (up to 2.0 atm) did not impact the lag phase (~12 h) (Fig. 2C and D). Although H2 was the limiting growth factor (Fig. S3), applying a PH2 higher than 0.70 atm reduced the optical densities by ~25% (Fig. 2C). In addition, elevated nitrogen partial pressures did not influence the attained optical densities (Fig. 2D), suggesting that C. necator cells can survive under high nitrogen pressures without impacting autotrophic growth. It should be noted that the gas composition was not monitored in the middle of the experiments because any gas extraction would have significantly changed the partial pressure inside the vials.

Substrate inhibition for heterotrophic growth.

C. necator growth under heterotrophic conditions was examined with acetate, glucose, glycerol, and mixed organic substrates (Fig. 3). Microbial growth was fast with a short lag phase (<6 h) at 1 to 2.50 g/L of acetate (Fig. 3A); however, elevated acetate concentrations (≥5.0 g/L) resulted in extended lag times (24 to 48 h) (Fig. 3A). The microbial growth of C. necator on acetate was previously reported to be severely inhibited at acetate concentrations greater than 1.0 g/L (9, 21, 33). It should be noted that the pH of the acetate solutions ranged between 6.0 and 6.8 at the end of the experiments (Fig. S4), indicating that pH did not significantly suppress the microbial growth at elevated acetate concentrations (19).
FIG 3 Heterotrophic growth of C. necator cells on acetate (A), mixed substrates (i.e., 33.3% acetate plus 33.3% glucose plus 33.3% glycerol) (B), glycerol (C), and glucose (D) (all experiments were conducted in duplicate [n = 2]). In the control vials, the growth solution with 1.0 g/L of organic substrate was used without microbial seed.
Along with acetate, C. necator cells were successfully cultivated on glucose, glycerol, and mixed substrates (Fig. 3B to D). Some studies have suggested that wild strains of C. necator cannot grow on glucose (34, 35). However, in the current work, C. necator Makkar and Casida 17697 was found to grow efficiently on glucose (Fig. 3D) or a mixture of glucose and other organic substrates (Fig. 3B), which matches other reports in the literature (36, 37).
In addition, the lag phase lengths (<12 h) were not impacted at 1.0 to 10.0 g/L of mixed substrates (Fig. 3B), glycerol (Fig. 3C), or glucose (Fig. 3D). However, the increases in the optical densities were not consistent with the increase in substrate concentrations. For instance, the optical densities at 10.0 g/L of organic substrates were only 4% to 40% higher than those at 1.0 g/L. According to these results, lower concentrations (≤2.50 g/L) of the studied organic substrate are recommended for efficient heterotrophic growth (9). It should be noted that the lowest pH at the end of the experiment was 5.26 for 10.0 g/L glucose (Fig. S4); however, the lag phase was not impacted by elevated glucose concentrations (Fig. 3D). These results suggest that lower pH did not impact the lag phase of C. necator when the bacterium was cultivated under heterotrophic conditions.

Organic substrates for mixotrophic growth.

Mixotrophic growth (Fig. 4) was more rapid than individual heterotrophic (Fig. 3) or autotrophic growth (Fig. 1) with similar substrate and gas conditions. The lag phases during mixotrophic growth on acetate and glucose (~5 h) were shorter than those on glycerol or mixed substrates (~12 h) (Fig. 4). Furthermore, the mixotrophic optical densities were greater than the individual heterotrophic or autotrophic ones. For instance, the highest OD with 1.0 g/L of mixed substrates under mixotrophic (1.23; Fig. 4) was greater than that under heterotrophic (0.77; Fig. 3B) or autotrophic conditions (0.78; Fig. 1). Previous reports on C. necator mixotrophic growth found that the microbial fixation of CO2 during heterotrophic growth needs an additional source of energy, such as H2, to improve the overall biomass yield (38) These results suggest that C. necator can utilize acetate, glucose, or glycerol as the main carbon and energy source and CO2 and H2 as additional carbon and energy sources (3, 9, 28).
FIG 4 C. necator mixotrophic growth on 1.0 g/L of acetate, glucose, glycerol, or mixed substrates (0.33 g/L acetate plus 0.33 g/L glucose plus 0.33 g/L glycerol). The headspace was replaced with the standard gas mixture (H2/O2/CO2 = 7:2:1) (all experiments were conducted in duplicate [n = 2]).

Hysteresis in two-stage cultivation.

C. necator strains can change metabolic pathways from heterotrophic to autotrophic/mixotrophic conditions very rapidly, as it takes less than 12 h for the complete transition (Fig. 5A to C). Consistently high optical densities (1.17 to 1.24) were attained when the heterotrophic conditions were switched to mixotrophic conditions at different substrate conditions (i.e., with existing acetate [Fig. 5A], after acetate consumption [Fig. 5B], or under starvation conditions [Fig. 5C]).
FIG 5 C. necator growth in two-stage cultivation systems. (A to C) The bacterium was first grown aerobically on 1.0 g/L acetate before switching to mixo- or autotrophic conditions after 12 h (A), 24 h (B), or 36 h (C) (all experiments were conducted in duplicate [n = 2]). (D to F) C. necator cells were first cultivated autotrophically (H2/O2/CO2 = 7:2:1) before switching to other conditions after 12 h (D), 24 h (E), or 36 h (F) (n = 2).
In addition, the transition from autotrophic to heterotrophic/mixotrophic growth was also rapid, taking 12 h or less (Fig. 5D to F). Similarly, the transition occurred at three different growth stages (i.e., during the exponential phase [Fig. 5D], at the end of the exponential phase [Fig. 5E], and after the steady-state phase [Fig. 5F]). However, a higher optical density was observed when the second stage was initiated after the end of the exponential phase (>24 h). According to our results, the two-stage cultivation system with initial heterotrophic growth followed by mixotrophic or autotrophic conditions can achieve higher optical densities in a short cultivation time (<24 h).


The impacts of cultivation conditions on the metabolic responses of C. necator were investigated. Under autotrophic conditions, the lag phases were short (<12 h) when a H2/O2/CO2 ratio of 7:2:1 was used. When the partial pressure of O2 was greater than 0.70 atm, the autotrophic growth was completely inhibited. Similarly, increasing the PCO2 value above 0.6 atm resulted in longer lag phases (>16 h) and severely impacted the optical densities. The elevated partial pressures of hydrogen (up to 2.7 atm) or nitrogen (up to 2.0 atm) had no impact on the lag phase lengths. Under heterotrophic conditions, the lag phase lengths (<12 h) were not affected at 1.0 to 10.0 g/L of glucose, glycerol, or mixed substrates; however, acetate concentrations of ≥5.0 g/L resulted in longer lag periods (>24 h). The concentration of organic substrates is recommended to be in the lower range (≤5.0 g/L) to decrease the lag phase time. With dual substrates, the mixotrophic optical densities were always better than the individual autotrophic or heterotrophic optical densities for the same substrate conditions. In two-stage cultivation systems, C. necator cells could completely switch among the heterotrophic, autotrophic, and mixotrophic pathways in less than 12 h. The highest optical densities (1.17 to 1.24) were achieved in two-stage systems with initial heterotrophic conditions, followed by mixotrophic or autotrophic conditions. The current results can improve the application of C. necator in large-scale industrial systems for biological CO2 fixation. In future work, the microbial consumption of organic and gaseous substrates during the experiments’ run needs to be measured. The impact of supplying high concentrations of organic substrates (>10.0 g/L) should also be studied. Moreover, the impacts of metabolic conditions on PHA accumulation by C. necator need to be investigated.


Bacterial strain.

C. necator Makkar and Casida 17697 was purchased from the American Type Culture Collection (ATCC) (Cedarlane Corp., Burlington, Canada). The bacterium was revived on solid agar medium and subsequently cultured in liquid media (see “Growth solutions and organic substrates,” below) according to the supplier’s instructions. For cell revival, the agar medium was prepared using 3.0 g/L beef extract, 5.0 g/L soybean peptone, and 15.0 g/L agar. After microbial revival, C. necator cells were continuously subcultured under autotrophic conditions (using the autotrophic medium [see “Growth solutions and organic substrates”] and a mixture of H2, O2, and CO2 at a ratio of 7:2:1 [Table 1]). In all experiments, a small amount of C. necator cells (~0.1 mL) from autotrophic subcultures was injected using sterilized syringes into the vials for the autotrophic, heterotrophic, mixotrophic, and transition experiments.
TABLE 1 Summary of the autotrophic gas mixture compositions applied in this study
ExptPartial pressure (atm) in the vial headspace
Preculture, autotrophic, mixotrophic, and transition experiments0.700.200.100
Inhibition by gaseous substrates0.700.20–2.200.100

Growth solutions and organic substrates.

The autotrophic growth solution included 2.30 g/L KH2PO4, 2.9 g/L Na2HPO4·2H2O, 1.0 g/L NaHCO3, 0.30 g/L NH4Cl, and trace amounts of minerals and vitamins (39). The initial pH of the autotrophic growth solution was 7.4 (SevenMulti pH meter; Mettler-Toledo International Inc., OH). For the heterotrophic and mixotrophic growth experiments, the same autotrophic growth solution composition was used, in addition to acetate (provided as CH3COONa), glucose, glycerol, or mixed substrates (33.3% acetate plus 33.3% glucose plus 33.3% glycerol [by mass]) at various concentrations (1 to 10 g/L). The initial pH of the heterotrophic growth solutions was neutral (between 7.2 and 7.6) for all organic substrate conditions. It should be noted that the four substrates have the same carbon load for the same concentration (e.g., 1.0 g/L of acetate, glucose, glycerol, or mixed substrates has 33 to 34 mmol/L C). For the autotrophic growth conditions, no organic substrate was added in the experiments.
All experiments (including the preculturing experiments) were conducted at 30°C in 17-mL Hungate glass vials with a 14-mL headspace volume and a 3-mL solution volume, unless mentioned otherwise. The vials were placed horizontally in a mechanical shaker (150 rpm) to increase the contact surface area between the medium solution and headspace gas. The growth solution and empty vials were autoclaved at 121°C for 20 min before the experiments. Then, the autoclaved solutions were aliquoted into the vials under the fume hood before injecting the microbial inoculum and gas composition (if required). All experiments were conducted in duplicate for the same conditions (n = 2). The measurement error (i.e., error bars) was calculated using the standard deviation formula.

Gas composition in the headspace.

For the autotrophic and mixotrophic experiments, the headspace air was replaced (using gastight syringes) with a gas mixture containing H2 (0.70 atm), O2 (0.20 atm), and CO2 (0.10 atm) (Table 1) (4, 16, 19). In the inhibition experiments using gaseous substrates, we examined the impact of various partial pressures of H2, O2, CO2, and N2 individually on C. necator autotrophic growth (Table 1). In the heterotrophic experiments, the headspace volume contained air only.

Transition between autotrophic and heterotrophic growth conditions.

The transition between heterotrophic and autotrophic conditions was examined in this study. For the transition from heterotrophic to autotrophic growth, C. necator cells were initially cultivated under heterotrophic conditions (1.0 g/L of acetate) before 1.0 atm of gas with the standard composition was injected at 12, 24, or 36 h (Table 1). For the transition from autotrophic to heterotrophic growth conditions, the experiment was started identically to the autotrophic growth experiment (Table 1), and 1.0 g/L acetate was injected at 12, 24, or 36 h.

Experimental analyses.

Microbial growth was monitored by measuring the optical density at 600 nm (OD600) (DR3000 spectrophotometer; Hach, USA). The biomass concentration was determined by measuring the net total chemical oxygen demand (COD) (Method 8000; Hach). The pH was measured using a SevenMulti pH meter (Mettler-Toledo International Inc.).
Samples were collected at the end of the experiments and tested for contamination using 16S rRNA gene Sanger sequencing, which was performed using the colony PCR procedure as previously described (40). In brief, the PCR was conducted with a pair of universal bacterial primers, 8f and 1482r (41, 42). The PCR mixture included 1 μL of 10-mM deoxynucleoside triphosphates (dNTPs), 0.25 μL of Taq polymerase, 5 μL of DNA template, 5 μL of 10× PCR buffer, 1.5 μL of 50-mM MgCl2, 1 μL of 10 μM forward and reverse primers, and distilled water (dH2O) to a final volume of 50 μL. The PCR program was initiated with heating to 94°C for 2 s; 29 cycles of 94°C for 30 s, 56°C for 30 s, and 72°C for 1 min; followed by a final cycle at 72°C for 10 min. Then, Sanger sequencing was conducted on a 3730 DNA analyzer (Applied Biosystems, Foster City, CA, USA) at the Mobix Lab (McMaster University, Canada). The sequencing results were viewed and modified using the FinchTV program (Digital World Biology, Seattle, USA) (43). The obtained merged sequences were checked using the NCBI BLAST tool to identify the bacteria (44). For all samples collected during the experiments, C. necator was identified with more than 98% identity, indicating that there was no biological contamination in any of the experiments. Thus, the OD results reported in this work can be considered representative for the growth of C. necator.


This study was supported by the Natural Sciences and Engineering Research Council of Canada (Discovery Grant, RGPIN-2019-06747, and Discovery Accelerator Supplement, RGPAS-2019-00102) and the Ontario Ministry of Research and Innovation (Early Researcher Award, ER16-12-126, and Ontario Research Fund—Research Excellence, RE09-077).

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Information & Contributors


Published In

cover image Applied and Environmental Microbiology
Applied and Environmental Microbiology
Volume 89Number 228 February 2023
eLocator: e02007-22
Editor: Pablo Ivan Nikel, Danmarks Tekniske Universitet The Novo Nordisk Foundation Center for Biosustainability
PubMed: 36719244


Received: 19 December 2022
Accepted: 9 January 2023
Published online: 31 January 2023


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  1. Ralstonia eutropha
  2. PHA-accumulating bacteria
  3. lag phase
  4. inhibition by high substrate concentration
  5. mixotrophic growth



Abdelrahman Amer
Department of Civil Engineering, McMaster University, Hamilton, Ontario, Canada
Civil Engineering Department, Menoufia University, Shibin Al Kawm, Al Minufiyah, Egypt
Department of Civil Engineering, McMaster University, Hamilton, Ontario, Canada


Pablo Ivan Nikel
Danmarks Tekniske Universitet The Novo Nordisk Foundation Center for Biosustainability


The authors declare no conflict of interest.

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