By Controlling the Concentration of Nutrients Added Into a Chemostat Continuously

Chemostat

A chemostat is defined as a steady-state bioprocess, where a microbial culture is continuously supplied with nutrients at a fixed rate and concomitantly harvested to keep the culture volume constant.

From: Methods in Enzymology , 2019

Continuous Cultures (Chemostats)☆

J. Gijs Kuenen , in Encyclopedia of Microbiology (Fourth Edition), 2019

Continuous Cultivation in the Retentostat or Recycling Chemostat

The chemostat is recommended to cultivate microorganism at submaximal specific growth rates. The technique can be used to grow pure cultures of microorganisms, or to enrich for microorganisms at low growth rates, that is, low substrate concentrations. In practice dilutions rate between 0.005 and 1.0  h^{−   1} have been used with chemostats of 1–2   L working volume. However the chemostat clearly has its technical limits when it comes to very low rates of growth and/or substrate supply, since the low flow of nutrients will cause fluctuations in the limiting substrate in the first place due to the drop size of the incoming medium and also to the potential effect of wall growth. An excellent tool to circumvent this problem is the use of a recycling- or retention-chemostat, in which part of the biomass is retained in the culture by using a membrane that keeps the microorganisms in the chemostat, but filters out the spent medium. Examples and suitable mathematical models will be cited in the following sections.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780128012383024909

Limits to Biodiversity (Species Packing)

L.B. Slobodkin , in Encyclopedia of Biodiversity (Second Edition), 2001

Glossary

Chemostat

Apparatus for growing microorganisms in a continually replenished medium.

Community

A multispecific aggregation of organisms in a particular location that may interact with each other.

Competition

More than one species utilizing one or more common resources.

Ecological niche

The set of requirements that must be met if a particular species is to survive. It is sometimes used to mean the place in which those requirements are met.

Ecosystem

A region with more or less clear boundaries, which contains a particular set of species and may be characterized by some set of meteorological, climatological, and geochemical properties.

Invasive or alien species

Species that have recently colonized some geographic regions different from the one in which they were initially described.

Isocline

A line along which some property remains constant.

Multidimensional niche

A tempero-spatial region defined by meeting a set of different requirements for viability of a particular kind of organism.

Niche dimensions

Ranges of values of environmental measurements in an ecological niche. A range of temperatures, salinities, or oxygen concentrations may be niche dimensions for a population of fish.

Packing

The placement of objects in a container.

Reification

The assignment of empirical reality to the referenda of a word or theory, regardless of the existence of any such referenda.

Species diversity

The number of different species in some area of interest. Global species diversity refers to all species. Local species diversity refers to some geographic region such as Hawaii or New York City.

Species packing

The study of how species on the same trophic level coexist in a limited region or container.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780123847195000885

Engineering Fundamentals of Biotechnology

P. Žnidaršič-Plazl , I. Plazl , in Comprehensive Biotechnology (Second Edition), 2011

2.21.3.4 Miniature Bioreactors

Miniature bioreactors with integrated sensors, usually termed microbioreactors, combine the small volumes of microtiter plates with the monitoring and control features found in bench-scale systems. The characterization of the engineering environment in miniature bioreactors has proved to be less complex than similar endeavor for shaken vessels, at least down to the milliliter scale. Most of the thorough work for the characterization of hydrodynamics and mass transfer in bench- and larger-scale-stirred vessels, and concomitant predictive correlations, apparently hold in miniaturized (milliliter volume) reactors. Compared to other miniature fermentation systems, scaling down from stirred tank reactors is less complicated, due to the inherent geometric similarity. Similar flow behavior and shear forces are therefore present in both systems, which make these devices an ideal tool for bioprocess development and optimization [15].

Pioneer work on the development of a miniature bioreactor for continuous culture was reported in 1994 by researchers from Space Biology Group of Swiss Federal Institute of Technology from Zürich, who realized a magnetically stirred 3   ml bioreactor with integrated pH, redox potential, and temperature microelectronic sensors for growing yeast cell cultures in space. Fresh medium could be fed to the incubation chamber with a micropump, while the inspection of the culture status was possible through the window and samples could be drawn through a silicone rubber septum [21]. Since then, several miniature bioreactors have been developed differing in vessel volume, mixing mechanism, aeration process, and monitoring and control levels, which were reviewed in recent publications [5, 14, 15, 18, 22].

Typical microbioreactor prototypes are realized in PDMS and PMMA by micromachining and multilayer thermal compression-bonding procedures with integrated online optical measurements for optical density, dissolved oxygen, and pH. Microfluidic connectors and fabricated polymer micro-optical lenses/connectors are integrated in the microbioreactors for fast setup and easy operation, and a magnetic stir bar is used for active mixing inside the reactor. The culturing of microbial cells in 150-µl-volume bioreactors in batch, continuous, and fed-batch operations has been demonstrated [22].

The key requirements to obtain a successful fermentation experiment in microbioreactor include: an adequate temperature and supply of nutrients, sufficient mixing, and a sufficient supply of oxygen for aerobic fermentations. Therefore, adequate mass and heat transfer are required. One of the most important operations in a submerged cultivation is mixing, which could be optimized as a compromise between ensuring homogeneous conditions and efficient mass and heat transfer and avoiding damage to the cells, and thus affects the quality of the achieved growth [5]. To achieve the active mixing, many microbioreactors include a stirrer bar mounted on and revolving around a rigid vertical post. Another active mixing approach involves moving the boundaries of the reactor. Periodic inflation of a series of air cushions in the ceiling of the reactor chamber created peristaltic movement of the liquid in the reactor. These methods, however, require delicate manufacturing, which result in higher production costs. Mixing in microfluidic bioreactors can also be achieved by passive methods that have no moving parts. Passive mixing, which requires some kind of flow of the broth through a microchannel, can also be subdivided into molecular diffusion and chaotic advection. Molecular diffusion can be enhanced with, for example, parallel lamination, while the chaotic advection further enhances mixing by stretching, folding, and breaking up the fluid streams. Although passive mixing elegantly removes the need for some kind of active mixing in the chamber, it creates additional fluidic requirements, such as pumps and possibly valves. Also, as with active mixing, great care must be taken to prevent dead zones where particles could sediment. Therefore, the future studies will focus on search for more simple and cheap mixing solution that provides sufficient mixing for a wide range of microorganisms [5].

An important part of aerobic fermentations is the supply of adequate aeration to the fermentation broth. A thin PDMS membrane is most commonly used for this purpose, as it allows diffusion of both oxygen as well as off-gases at sufficiently high rates while additionally ensuring continuous sterility. In microbioreactors with a bubble-free system, aeration can be achieved for example by integrating a gas-permeable membrane into the top side of the microbioreactor chamber. Despite high surface-to-volume ratios of microsystems and therefore a large interfacial area, efficient transport of oxygen-rich fermentation broth away from the membrane is also important, and can be achieved by providing sufficient mixing. Another method of achieving sufficient oxygen supply is to increase the oxygen content in the gas phase. The dissolved oxygen concentration in microbioreactors is typically measured by optical sensors ( Figure 5 ) [22] with the use of fluorescence sensor spots, based on the quenching of fluorescence by oxygen. The optical sensors can be manufactured in small sizes, are easy to integrate, insensitive to ambient light, relatively cheap, and nonreactive. The dissolved oxygen concentrations in microbioreactors can also be measured by an amperometric biosensor, ultra-microelectrode array (UMEA), which measures oxygen concentration based on the electrochemical reduction of oxygen.

Figure 5. Illustration of the microbioreactor setup with indicated optical fibers (dashed lines) and electronic wires and fluid tubes (solid lines).

Reproduced from Zhang Z, Perozziello G, Boccazzi P, et al. (2007) Microbioreactors for bioprocess development. Journal of the Association for Laboratory Automation, 12: 143–151. Copyright (2007), with permission from Elsevier.

The most commonly applied miniature pH sensors for use in microbioreactors are optical sensors based on fluorescence sensor spots ( Figure 5 ), also called optodes, and solid-state, ion-sensitive, field-effect transistor (ISFET) pH sensor chips. Despite some limitations, both sensors have been shown to provide rapid and precise pH measurements over a long period of time. While a real-time pH measurement in microbioreactors is fully feasible, pH control in microbioreactors is still in the development phase. Online monitoring of the cell density in microbioreactors is normally realized via optical probes, so that the light from light-emitting diodes is guided into the microbioreactors with optical fibers, sent through the reactor chamber, and then guided to a photodetector ( Figure 5 ). Though optical density (OD) measurement via optical probes has proved to be the most feasible way of estimating cell density in microbioreactors, gas bubbles in the reactor chamber can interfere with OD measurements. In addition, the OD measurement measures the total cell concentration (viable and death cells). Cell density of live cells can only be estimated in microbioreactors by means of impedance spectroscopy. This method applies an alternating current (AC) electrical field to the culture (only cells with intact membranes are able to polarize charge) and measures the cell conductivity as a function of frequency [5].

External syringes or peristaltic pumps are generally used to pump fresh culture medium into the reactor ( Figure 5 ), which then also passively (due to the fixed reactor volume) pushes the same amount of culture broth out of the reactor in the case of a continuous reactor with constant volume. Based on this design, a multiplexed system for parallel operation of four microbioreactors was reported [23].

A membrane aerated hollow-fiber microbioreactor was also developed which consists of an acrylic glass module equipped with two different types of membrane fibers for substrate and oxygen supply. Online measurement of dissolved oxygen and optical density was also integrated into the bioreactor. Due to very efficient oxygen transfer during cultivation of model microorganism, better cell growth than in shaking flask experiments was achieved. The potential of this microbioreactor as a a screening tool for a wide range of microorganisms and cells was reported and future improvements including better monitoring (pH, CO₂), as well as continuous culture assay and integrated downstreaming is predicted [24].

2.21.3.4.1 Microchemostats

Chemostats are continuously operated bioreactors where growing cells reach a steady state condition at which specific growth rate, as well as biomass, substrate and the product concentrations remain constant. They are therefore an extremely powerful tool for the precise analysis of cell metabolism and with the miniaturization their role in biological and physiological research, and for the growth model parameters evaluation, got even higher potential.

One of the first continuously operated microbioreactor had six units with a working volume of 16 nL working in parallel on a single chip. One of the main advantages enabling to monitor the programmed behavior of bacterial populations for hundreds of hours was an active approach to preventing biofilm formation. The microchemostat, where mixing was obtained by circulating flow in a microfluidic loop, operated by alternating continuous circulation with dilution and cleaning by means of a lysis buffer. The miniaturized device enabled automated culturing and monitoring of populations of 100 to 10⁴ bacteria with instantaneous single-cell resolution [25].

Another version of the microchemostat was an improved version of a microbioreactor presented in Figure 5 . The fouling caused by biofilm formation was addressed by the coating method using copolymer films on PMMA and PDMS layers. Furthermore, the problem of chemotaxial back growth of bacterial cells into the feed line was solved by local heating. The microbioreactor was integrated with optical density, pH and dissolved oxygen real-time measurements, which confirmed steady state conditions [26].

However, it is still not fully validated whether microchemostats really maintain continuous culture with the precision of the conventional bioreactors. Furthermore, there is a need to design microchemostats for extremofiles growing at e.g. high pressure, high temperature or at extreme pH conditions, and for long-term monitoring for exceptionally slow growth states, which would open up the study of relatively intractable microorganisms [14].

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780080885049001021

How Fast Can We Grow?

Alexei Vazquez , in Overflow Metabolism, 2018

4.5 The Chemostat Experiment

The chemostat is an experimental apparatus where the chemical environment can be maintained static and nutrient availability can be controlled by the experimenter. This is achieved by culturing cells in a vessel subject to continuous supply of nutrients and continuous overflow of content exceeding the vessel volume (a glass under a dripping tap is good analogy). The chemical environment in the chemostat vessel is determined by the chemical composition of the feeding medium, the dilution rate (rate of liquid volume addition relative to the vessel volume) and the type of cells cultured. After some transient period, the proliferation rate of the cells in the vessel will match the dilution rate [89], provided that the cells remain in suspension and the dilution rate does not exceed the maximum proliferation rate.

Cells need sources of carbon, nitrogen and other elements to proliferate. The relative abundance of these sources in the feeding medium determines which nutrient is the limiting factor for growth. For every chemostat experiment we need to specify the limiting element: carbon, nitrogen, etc. It is also common to name the experiment after the specific source of limiting element. For example, in a glucose-limited experiment, carbon is the limiting element and glucose is the carbon source. We also need to specify whether the experiment was conducted in the presence (aerobic) or absence (anaerobic) of oxygen. The chemostat is the experimental apparatus of choice to investigate the metabolism of cells that can grow in suspension at different proliferation rates. Since proliferation rate is in this context a measure of metabolic rate, we can use the chemostat to investigate the differential utilization of oxidative phosphorylation and fermentation at different metabolic rates.

Fig. 4.1 shows the typical output of a chemostat experiment for a glucose-limited culture of yeast cells under aerobic conditions, based on data reported in reference [90]. At low dilution rates there is no fermentation. I will call this scenario the Pasteur phase: 'oxidative phosphorylation represses glycolysis'. However, after a threshold dilution rate fermentation kicks in and its rate increases linearly with increasing the dilution rate. I will call this scenario the Brown–Warburg–Crabtree (BWC) phase, where aerobic fermentation is manifested.

Figure 4.1. Ethanol excretion rate by yeast cells. The symbols represent measurements in units of mol ethanol/g dry weight/h [90], multiplied by the typical biomass density of a cell 0.3   kg/L. The line represents the theoretical prediction from Eqs. (4.24) and (4.25).

The transition from pure oxidative phosphorylation in the Pasteur phase to mixed oxidative phosphorylation with fermentation in the BWC phase is a consequence of two simple facts pointed in the previous section. Oxidative phosphorylation has a higher yield of ATP per molecule of glucose than fermentation but fermentation can sustain a faster proliferation rate. When the proliferation rate exceeds the maximum growth rate allowed by oxidative phosphorylation (μ _max,O ) fermentation becomes obligatory.

To model the concomitant occurrence of aerobic and fermentation metabolism we extend the cell growth model considered above after dividing the energy production by the corresponding components associated to oxidative phosphorylation and fermentation. This result in the updated volume fractions balance

(4.20) $\varphi ={\varphi }_0+{\varphi }_R+{\varphi }_F+{\varphi }_O$

where ϕ _F and ϕ _O are the cell volume fractions occupied by the fermentation and oxidative phosphorylation machinery, respectively. From postulate (1), we can also write the balance of protein content

(4.21) $P=P_0+n_F{\varphi }_F+n_O{\varphi }_O+n_R{\varphi }_R$

where n _F is the number of amino acids per unit of molar volume of fermentation enzymes and n _O is the number of amino acids per unit of molar volume of the oxidative phosphorylation machinery.

From postulates (2)–(4), we can write the metabolic balances, of protein synthesis/growth

(4.22) ${\varphi }_R{h}_R=\mu P$

and energy production/consumption

(4.23) $h_F{\varphi }_F+h_O{\varphi }_O=e_P\mu P$

where h _F and h _O are the horsepower for energy generation by fermentation and oxidative phosphorylation and e _P is energy cost of protein synthesis per unit of amino acid.

Using Eqs. (4.20)–(4.23), we can determine the combinations of aerobic and fermentation metabolism that can sustain the energy demand of cell growth as a function of the growth rate (Fig. 4.2). The solutions go from all energy being generated by fermentation (assuming that there is enough sugar to do so) to the solution with minimum fermentation. We can estimate the minimum fermentation rate that is required to sustain growth – r _F,min(μ) – by interpolating between the lack of absolute fermentation requirement at μ _max,O and the absolute requirement at μ _max,F

Figure 4.2. Combinations of aerobic and fermentation metabolism satisfying the energy demand at different growth rates.

(4.24) $r_{F,\min }\left(\mu \right)=\{\begin{array}{ll}0\hfill & \mu \ge {\mu }_{\max ,O}\hfill \\ \frac{\mu -{\mu }_{\max ,O}}{{\mu }_{\max ,F}-{\mu }_{\max ,O}}{r}_{F,\max }\hfill & {\mu }_{\max ,O}<\mu \le {\mu }_{\max ,F}\hfill \end{array}$

where r _F,max is the maximum fermentation rate at the maximum growth rate sustained by fermentation. r _F,max is obtained from Eqs. (4.20)–(4.23) setting ϕ _O =0 and μ=μ _max,F resulting in

(4.25) $r_{F,\max }=\frac{e_P{\mu }_{\max ,F}\left(P_0+n_R\left({\varphi }_{\max }-{\varphi }_0\right)\right)}{1+e_P{\mu }_{\max ,F}\left(n_R-n_F\right)/h_F}$

Using the parameter estimates for yeast cells (Table 4.3) and Eqs. (4.24)–(4.25), we obtain the theoretical line depicted in Fig. 4.2. It clearly captures the qualitative behaviour of the experimental data. No fermentation below the threshold growth rate and a linear increase of fermentation above the threshold. The onset of overflow metabolism is basically the maximum growth rate that can be sustained by oxidative phosphorylation (μ _max,O ). The theoretical estimate is slightly higher than what observed. Yet, it is quite a good estimate given that we only took into consideration the protein requirements of cell growth. Addition of other cell components, particularly lipids, will certainly reduce the theoretical estimate of the maximum growth rate that can be sustained by oxidative phosphorylation.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780128122082000044

Agricultural and Related Biotechnologies

C.G. Trick , in Comprehensive Biotechnology (Second Edition), 2011

4.24.1 Introduction

Microalgae are commonly considered to be one area of biotechnology that can provide societal benefits and low economic or ecological risks. The design and implementation of microalgae growth and harvesting technology will influence the biotechnology outcome. Of the commonly considered growth strategies are the large-scale pond enclosures, runway-based algal reactors, or the fermenter-type closed growth culture systems. Each of these systems is designed primarily to process an initially high level of artificial or wastewater nutrients into new algal biomass or to use the consumption of nutrients by algae in the reduction of nutrient levels in the wastewater [2] . A less utilized system is the flow-through continuous-culture growth reactor that regulates the physiological state of the cells but a strict control of the cellular growth rate and the composition of the media, particularly the level of the limiting nutrient. These continuous-culture reactors, termed chemostats, provide an alternative for the batch culture experiments and are particularly valuable in ecological studies, in microalgal selection and isolation, and in the production of secondary metabolites, where the efficiency of cellular production is balanced against the high cost of product generation. Chemostat systems are defined as a flow-continuous-culture system where the biomass of the cells is regulated by the composition of the limiting nutrient (compared with the cellular needs) and the growth rate of the cell is maintained through the dilution rate of the reactor. Thus, production of algal metabolites that are produced at rates dependent upon the limiting nutrient and the growth rate can be maximized in the experimental design. At the foundation of chemostat studies is the recognition that microalgae differ substantially in their nutrient needs (cell nutrient quota), their efficiency at obtaining nutrients from the media or environment (nutrient uptake kinetics), their efficiency of converting elements and energy into new biomass (growth kinetics), and the biochemical composition or elemental stoichiometry of the cells. Chemostat experiments or applications exploit the steady-state or balanced growth cellular model at the ecological, ecosystem, or physiological levels – creating an understanding of microalgal products not achieved through batch culture-based studies where the primary product is biomass accumulation (mass production of lipids, carbohydrates, and other structural cell components) or the removal of macronutrients into a particulate cellular fraction [11].

The value of the chemostat experimental setup is twofold:

1.

With some experimental modification, chemostats represent the common growth kinetics of natural water systems – enabling theexperimental modeling of key components of carbon and energy transfer, metabolite exchange, and competition for limiting resources. Algal communities that are competing for the same nutrient, scavenging the excess nutrient, and dominating the community by outgrowing the other algal competitors, dominate most ecosystems. There are three levels of experiments in this approach to the application of chemostats to natural ecological models. First, there are chemostats that apply the flow-through model to a natural complex community to assess the competitive success rates of members of the community under defined environmental conditions. Second, there are the traditional experiments where the cellular growth rates and cell nutrient quotas are assessed against a defined limiting condition. This approach provides cell-specific experimental evidence that is used in ecological models. Third, by further narrowing down the choice of host organism and applying low concentrations of the test solution or compound, this type of chemostat system shows promise as an effective ecotoxicity model.

2.

Chemostats are valuable in biotechnological applications where the production of individual compounds can be maintained at the highest level by modifying and defining growth conditions based on growth rate and the composition of the medium. Chemostat research in cellular biotechnology has evolved to not only include experimental setups that define the limiting nutrient, but also modify or define the not-limiting nutrients/light to enhance specific metabolite production.

There is a long history of the application of chemostat studies to ecological processes, and this has developed into a defined research in wastewater processing. There is a new line of research using chemostats to biodegrade and scavenge environmental contaminants. The application of chemostat principles and biotechnology is also undergoing a renaissance. The recognition that algal cells have specific, unique, or novel metabolites that can be enhanced under specific, but defined, growth conditions is relatively unexplored but shows promise, as proper-scale algal bioreactors using chemostat principles are designed.

Algal chemostats are experimental chambers where the growth state of algae can be maintained through the constant, primarily low, level of nutrients. This flow-through experimental apparatus maintains cells in steady-state conditions – with the composition of the cells constant from one day until the next. In general, the setup comprises a known algal medium placed into a well-mixed reactor and a pump that adds the medium at a fixed dilution rate. The reactor chamber is inoculated with a single algal culture, a mixed or natural algal culture, or a combination of algae with another competitor or predator. As the new medium enters the growth chamber and equal volume of media with cells are removed, ensuring that the volume of the chamber remains constant, the result is that the nutrient status of the cells in the chamber remains constant; cells that cannot compete with the rate of exchange of nutrients are removed and the cell nutrient conditions are constant.

Algal chemostats are best understood when compared with the batch-culture setup. In this more traditional setup, the inoculum of cells is provided with a generally rich medium, under suitable conditions for growth. As the cells grow, the nutrients are removed from the medium and the density of the cells increases. Over the time in the culture, the cells are exposed to a continually declining level of nutrients, an accumulation of metabolic waste products, and the decline in the amount of light available to the cells. Thus, it is difficult to relate the physiological or biochemical status of the cells to any specific environmental condition. Chemostats provide the constant physical and nutritional conditions that allow for a clear relationship between the environmental conditions and the cell physiology.

Since the initial description of the continuous-culture setup by Monod in 1950, the chemostat has drawn attention in photosynthesis studies, ecological and competition experiments, and in the regulation of cellular compounds. With the continued development of algal biotechnology, the chemostat growth system has attracted attention as a means to control and enhance individual chemical components (pigments, toxins, and oils).

Chemostats along with turbidostats are the two methods of continuous culture. Continuous culture indicates that the cells are harvested at some stage in the growth cycle, regulated by either the amount of limiting nutrient in the medium (chemostats) or the density of the cell biomass (turbidostats). The harvested cells, often referred to as cells in balanced growth, reflect a constructed nutrient environment, not a rapidly altering environment. In the case of the turbidostat, the cell density is maintained at an established level of cell suspension by diluting the medium with fresh, nutrient-rich medium as the cell density reaches the prescribed level. The small dilution of the culture with the fresh medium provides nutrients for growth and ecological space for the cells to divide before the culture is diluted again. Thus, cells from turbidostats are in steady state and are expelled with excess nutrients. These cells are not nutrient limited but are in steady state with respect to factors such as light, pH, and temperature.

By contrast, chemostat-generated steady-state conditions are defined by the element in the medium that is limiting – that is, the nutrient that is at the lowest available concentration compared with the needs of the cell. It is this limiting nutrient that defines the physiological status of the cells. For algal cultures, the modification of the limiting nutrient provides cells of different, but defined, physiological states [8]. For example, cells grown in media that contains low levels of nitrate will consume all the available nitrate, leaving behind in the cell medium the nutrients in excess, and express the phenotype of a nitrogen-limited cell. Other nutrients that commonly have been reduced in the chemostat medium include phosphate, silicate (for diatoms), iron, and vitamins. Thus, each setup creates a specific cell phenotype related to the desired environmental conditions. This provides a great diversity in the design of chemostat-based experiments when environmental conditions such as light, temperature, and pH can be studied in combination with the prescribed limiting nutrient.

While the limiting nutrient is paramount to the description of chemostat experiments, the rate at which fresh media is added establishes the population density and the growth rate of the cell culture. Thus, the steady-state situation is based on the supply of nutrient relative to the growth rate of the cells. Thus, the early chemostat experiments investigated the relationship between nutrient availability and achieved cell growth rates in a continuous-flow-through culture apparatus. This can be expressed as

$\text{The rate of change in cell density = growth of the cells - loss of cells through washout}$

By denoting the rate of growth as µ and the loss of cells as a function of the dilution rate (volume entering over the volume of the culture vessel   = D), the Monod model equation is

$\text{d}N/\text{d}t=N(\mu -D)$

where N is the number of cells in the culture vessel and t is the time (hours or days) [13]. From this equation, the steady-state condition will be achieved when µ  = D.

This steady-state equation can be related to the growth rate of the cell if the replacement rate of the nutrients into the cells is considered. This is done by comparing the growth rate with the Michaelis–Menten nutrient uptake rates, where

$\mu ={\mu }_{\max }(S/(K_{\text{s}}+S))$

In this situation, the maximum growth rate (µ _max) will be achieved when the concentration of the nutrient (S) is highest when compared with the half-saturation value (K _s) for the limiting nutrient. This equation defines that the dilution rate of the chemostat will set the growth rate of the population and the biomass of the chemostat will be set to the relative need of the limiting nutrient.

Studies accepted then challenged the Monod model of growth regulation. In the Monod model, the nutrient uptake rate is proportional to the reproductive rate [3]. This does not hold when cells have the capacity to store nutrients in the cell biomass. This stored nutrient, sometimes called cell quota, represents the minimum level of nutrients in the media that will support growth. The cell quota is not a constant but varies with the growth rate of the cell and the level of extracellular nutrient. The Monod expression can be modified to include the level of internal nutrient pool as defined by the Droop model (also referred to Caperon–Droop model) [6] where the concentration of the cellular nutrient pool is stated as Q. The resulting relationship is

$\mu ={{\mu }^{\prime }}_{\max }(1-k_{\text{q}}/Q)$

where k _q is the minimum level of limiting nutrient when the quota is the smallest and µ′_max is the growth rate when the nutrient is in great excess. Further modifications have indicated that by assuming that Q does not vary with the level of substrate, the Droop–Caperon equation can be simplified further without considerable loss in the understanding of the chemostat model [4]. Burnmaster's modification more directly links the rate of nutrient uptake (v) to the relative need of the cell for the limiting nutrient:

${{\mu }^{\prime }}_{\max }={\nu }_{\max }/(Q_{\max }-Q_{\min })$

Over the 60 years since introduced by Monod, balanced steady-state growth by chemostats has attracted considerable attention by competition theorists and mathematical ecologists. There is a rich level of discussion and theory concerning the detailed analysis of growth under these defined conditions and under variable steady-state scenarios. However, the reason for this high level of detail is that the physiological status of algal cells and cyanobacteria is directly related to environmental conditions and to their application in biotechnology. Concepts of nutrient limitation and growth rate are critical in the metabolic balance of the cells – and thus their biochemical composition.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780080885049002452

Microfluidics in Cell Biology Part B: Microfluidics in Single Cells

L.J. Holt , ... M. Delarue , in Methods in Cell Biology, 2018

5 Discussion

The proposed mechano-chemostats are extremely flexible microfluidic devices that enable the study of compressive mechanical stress. They are compatible with optical imaging, and the control of the chemical environment makes it possible to perform staining like FISH or immunofluorescence in situ. Moreover, high-throughput devices can be used to rapidly screen for multiple conditions, either genetic or chemical, that alter cell proliferation/survival under a compressive mechanical stress.

Even though our mechano-chemostats were developed for the fungus S. cerevisiae, we were also able to use them to study other yeasts including S. pombe and Candida albicans. The geometry of mechano-chemostats, notably the valve and the size of the nutrient channels, can either be downscaled to study smaller microbial organisms, or upscaled for the study of mammalian cells. The advantage of mechano-chemostats over existing techniques (Alessandri et al., 2013; Helmlinger et al., 1997; Minc et al., 2009; Mishra et al., 2017) resides in the versatility of their usage. They allow dynamic control of pressure, along with real-time control of the chemical environment. We believe that our approach could be adapted to ask a variety of fundamental questions regarding the impact of compressive stress on living organisms.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/S0091679X18300736

Growth Kinetics

Ghasem D. Najafpour , ... Ghasem Najafpour , in Biochemical Engineering and Biotechnology, 2007

5.6.8 Modified Chemostat

With cell recycling, chemostat efficiency is improved. To maintain a high cell density the cells in the outlet stream are recycled back to the fermentation vessel. Figure 5.10 represents a chemostat unit with a cell harvesting system. The separation unit is used for harvesting the cells and recycling then to the culture vessel to increase the cell concentration.

FIG. 5.10. Chemostat with a cell recycle stream.

The material balance in a constant volume chemostat is derived based on cell balance as shown in the following equations. Material balance in a chemostat with recycle, ρ _cell:

(5.6.8.1) $\frac{\text{d}X}{\text{d}t}=\left(\frac{F}{V}\right)[X_o-X\left(1+\tau \right)]+\mu X+\left(\frac{F}{V}\right)\cdot cX$

where τ = recycle ratio

c = the factor by which the outlet stream is concentrated before return

For steady-state,

(5.6.8.2) $\mu =D\left(1+\tau -\tau c\right)$

Multi-stages of continuous culture are designed to use the outlet of the first vessel as the inoculum for the next stage. If intermediate metabolites are used as feed for another microorganism, sequential continuous culture is useful. The dilution rate for each vessel may be different to the other vessel. It is also possible to supply different nutrients for each stage of fermentation vessel. It is common to operate earlier stages as aerobic and subsequent stages in an anaerobic condition. In addition, if unused substrate leaves the product stream, it can be used in the next stage even at low substrate concentration. The kinetic representation may show a slower rate and even drop to zero-order. Figure 5.11 shows two stages of a chemostat in operation.

FIG. 5.11. Two stages of CSTR fermentation vessels in series.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780444528452500057

Continuous Cultures (Chemostats)

J.G. Kuenen , O.J. Johnson , in Encyclopedia of Microbiology (Third Edition), 2009

Important Aspects of Continuous Culture

1.

Continuous culture in a chemostat enables the reproducible growth of bacteria and other microorganisms. Consequently, the chemostat is an appropriate tool for quantitative (eco)physiological research.

2.

Microorganisms can be studied while growing at submaximal rates.

3.

The effect of different growth limitations on the metabolism of the cells can be measured reproducibly.

4.

The chemostat is also appropriate for studying the competition of microorganisms and mutants for growth-limiting substrates.

5.

Continuous cultivation can also be performed under fluctuating environmental conditions.

6.

Recent publications show the great value of chemostat cultivation for functional genomic research and for the selection of industrially relevant mutants.

7.

The commercial availability of well-designed chemostat equipment greatly facilitates the introduction of continuous cultivation in the laboratory.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780123739445001127

Agricultural and Related Biotechnologies

E.D. Leonardos , B. Grodzinski , in Comprehensive Biotechnology (Second Edition), 2011

4.14.4.1 Greenhouses as Bioreactors for Vascular Plants

Just as algal chemostats (i.e., bioreactors) have been developed to produce aquatic photoautotrophs for commercial applications ( see Chapter 4.24), there have been improvements in the design of protected structures such as greenhouses and plant phytotrons for growing a range of vascular crops [16]. Although glasshouses have been in use for over a century, now the plant microclimate in a modern greenhouse is under computer control. The net result of the new technologies has been the growth of multibillion dollar industries in Europe (e.g., Holland) and North America (e.g., Canada). In the winter months, natural photoperiod and low temperatures obliterate outdoor agriculture, but consumer demands for fresh produce are year-round. It suffices here to say that CO₂ enrichment works to enhance photosynthesis and plant productivity. CO₂ enrichment is essential for commercial success, because plants are photoautotrophs and CO₂ is the primary substrate of photosynthesis. Greenhouses are essentially huge bell jars. When the sun is shining, the interior CO₂ levels of the greenhouse can be depleted markedly in the light ( Figure 5 , red line). In winter months, especially with vents closed to prevent heat losses, the depletion in CO₂ can be rapid and crop is in a severe photorespiratory condition. The canopy is operating near the CO₂ compensation point (CCP) and certainly suboptimally for RuBisCO. Briefly, CCP is the value of CO₂ at which there is no net C fixation, and photosynthetic C gain equals respiratory C losses. The CCP for C₃ leaves is normally between 40 and 60   ppm, whereas for C₄ photosynthetic types they are lower, 4–8   ppm. Whole-plant CCP is higher due to plant respiration and mutual shading. Greenhouse producers have known all this for decades [16] and taken appropriate remedial action by adding CO_2. The methods of CO₂ delivery to the canopy are varied. The two main protocols involve using bottled CO₂ or deriving the CO₂ from the off gases of combustion used to generate heat. Regardless of how the substrate for photosynthesis is delivered, it is under tight computer control and target levels for a crop are set based on researched response curves to varying irradiation levels, CO₂ responses, and temperature [22, 33]. However, it needs to be emphasized that in setting these environmental variables, they are not done alone. All other plant inputs such as nutrients and H₂O delivery have also been optimized for latitude and seasonal production needs. Production protocols for managing greenhouses are highly evolved, and photoperiod control, artificial lighting, and temperature are used to match as much as can be accomplished economically [16] the photoperiod requirements of flowering and fruiting crops are matched to optimize production year-round (see Chapter 4.17). Today, the commercial trend is toward geothermal heating and cogeneration systems to reduce heating costs associated with fossil fuels. But regardless of how cost-effective inputs might become, CO₂ enrichment to levels around a 1000   ±   200   ppm are normal.

Figure 5. Daily patterns of atmospheric CO₂ concentrations experienced by plants in field and controlled environments.

The benefits of an enhanced rate of plant photosynthesis can be many, including increased final yields and higher quality of produce. A shortened period of plant growth to harvest maturity in a greenhouse can provide significant economic returns, such as reducing heating and maintenance costs. Timing of crop production schedules is virtually ignored in other discussions of CO₂ enrichment for crops, because researchers are focused on seasonal field crops and final yield.

Where does this position us for the future as we already use very advanced greenhouse systems for selected fresh produce (e.g., vegetables, herbals, and ornamentals). One possibility is the adaptation of greenhouse technologies for vertical farming in tall buildings and in mine shafts where heating is minimized because of geothermal warming. Closed structures offer intriguing possibilities for expanding the way we approach the pressure of increasing population and reduced land use for agricultural food and bioproduct production (see Chapters 4.01 and 4.02). One idea is to grow transgenic plants with novel bioproducts, such as pharmaceuticals, in sealed environments rather than in the field. Closed environments offer the possibility of fidelity in the control of key environmental variables such as lighting, temperature, CO₂ levels, nutrients, and water recycling. The outdoor or classical open-field production of these plants is restricted in many countries by legislation governing production of genetically modified organisms.

A somewhat esoteric spinoff of this technology is for long-term manned exploration of space where vascular plants as well as algal chemostats would generate O₂ as well as scrub CO₂. Higher plants through evapotranspiration generate potable water and CO₂ enrichment would benefit production overall [14]. Regardless of the perceived sanity or insanity of these extraterrestrial applications, knowledge about photosynthesis, semi-closed environments, and production in closed environments is important. Interestingly, both herbaceous and woody plants are incorporated by architects as part of aesthetically pleasing, indoor, air-purification systems (see Chapter 4.26).

In predicting the viability of the greenhouse industries what seems to threaten their further expansion, is globalization and the rapid movement to market of fresh produce. In most parts of the world, fresh produce of all kinds are shipped out of season over long distance. Improvements in our understanding of post-harvest problems (see Chapters 4.27 and 4.28) and aggressive international marketing mean that imported fresh produce of high quality is available year-round at reasonable prices. It is not clear if established operations can remain profitable, but given resilience and innovation of this industry, and the rising costs of jet fuel, it seems reasonable to predict that local production or at least their technologies will flourish.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780080885049002415

CLOSTRIDIUM | Clostridium Acetobutylicum

Hanno Biebl , in Encyclopedia of Food Microbiology, 1999

Development of Fermentation and Product Recovery

Continuous fermentation in a chemostat mode has proved to be an effective tool to increase productivity in the butanol–acetone fermentation. Phosphate is an appropriate limiting factor but cultivation without nutrient limitation is also possible as the accumulating products limit growth and give rise to steady states. Usually lower product concentrations are obtained than in batch culture but by application of two stages, an acid-forming growth stage at high dilution rate and a solvent-forming fermentation stage at low dilution rate, a solvent concentration was achieved approaching the usual batch concentration of 20  g l⁻¹ solvents.

To increase the relatively low productivity of chemostat cultures (0.5–2   kg solvents/(h m³)) two techniques, both designed to operate at elevated cell densities, were studied. With cell immobilization, spores are entrapped in gel beads or attached to solid particles using a low-growth medium, which is preferably nitrogen limited. Calcium-alginate beads and beechwood shavings have been successfully tested. Cell recycling involves permanent withdrawal of cell-free culture liquid into an external filtration unit and returning of the more concentrated culture to the fermenter. With both methods a productivity increase of about fourfold was achieved in comparison to the free-cell continuous culture. The rates obtained vary according to the amount of added complex substances such as yeast extract and peptone, the maximum being at 3   kg/(h m³).

The low final solvent concentration attained in the butanol–acetone fermentation and the high energy requirement for distillation of butanol, the boiling point of which is greater than that of water, has initiated a search for alternative solvent recovery processes. The main emphasis was put on product removal procedures that are integrated in the fermentation and thus increase productivity by reducing the concentration of toxic products in the culture.

As suggested above in relation to the industrial production, extraction by a water-immiscible liquid in direct contact with the culture has the advantage of being simple to realize. Good results have been obtained with oleyl alcohol, diluted with decane to reduce viscosity. Octanol has also proved to be a useful extractant, but as this compound is slightly toxic to the clostridia, it was necessary to separate the cells from the culture liquid by microfiltration. The solvents are extracted selectively and can be recovered by distillation at a relatively low energy input. A modification of the liquid–liquid extraction is known as perstraction, the culture being separated from the extractant by a solvent-permeable membrane. It avoids formation of emulsions between the phases, and the extractant need not be sterilized and cannot affect the culture.

Inert gas is used to remove the solvents in variants with and without membranes. Gas-stripping, i.e. direct sparging of gas through the fermenter, is likewise attractive because of its simplicity. The microorganisms are not affected, and the products are easily recovered by condensation, with less energy consumption than with distillation of liquid extractants. It has been suggested that the self-produced fermentation gases, carbon dioxide and hydrogen, are used instead of expensive nitrogen. The membrane modification of gas-stripping, pervaporation, requires an extended tubing system which is immersed in the fermentation vessel. The solvents evaporate through the membrane and are drawn off by vacuum or sweep-gas. As the available membranes only allow passage of the solvents the acids accumulate in the culture and may stop the fermentation. This problem was solved by low-acid mutants that were able to reutilize all of the acids.

Adsorption to solid materials such as silicalite or polyvinylpyridine has also been tested. Relatively low loading capacity, high estimated costs for the adsorbants and the heat for desorption of the solvents presently diminish the chances for this method. For external application reversed osmosis has been evaluated and found to be more favourable than distillation.

Generally speaking the in situ recovery methods are interesting, but require high capital expenditure and permanent monitoring by the operator, and although their technical feasibility is established they need further development at the engineering level.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B0122270703003858

greenethilbod.blogspot.com

Source: https://www.sciencedirect.com/topics/immunology-and-microbiology/chemostat

Share this post