*Article* **Determination of Dissolved CO2 Concentration in Culture Media: Evaluation of pH Value and Mathematical Data**

#### **Amir Izzuddin Adnan 1, Mei Yin Ong 1, Saifuddin Nomanbhay 1,\* and Pau Loke Show <sup>2</sup>**


Received: 23 September 2020; Accepted: 26 October 2020; Published: 29 October 2020

**Abstract:** Carbon dioxide is the most influential gas in greenhouse gasses and its amount in the atmosphere reached 412 μmol/mol in August 2020, which increased rapidly, by 48%, from preindustrial levels. A brand-new chemical industry, namely organic chemistry and catalysis science, must be developed with carbon dioxide (CO2) as the source of carbon. Nowadays, many techniques are available for controlling and removing carbon dioxide in different chemical processes. Since the utilization of CO2 as feedstock for a chemical commodity is of relevance today, this study will focus on how to increase CO2 solubility in culture media used for growing microbes. In this work, the CO2 solubility in a different medium was investigated. Sodium hydroxide (NaOH) and monoethanolamine (MEA) were added to the culture media (3.0 g/L dipotassium phosphate (K2HPO4), 0.2 g/L magnesium chloride (MgCl2), 0.2 g/L calcium chloride (CaCl2), and 1.0 g/L sodium chloride (NaCl)) for growing microbes in order to observe the difference in CO2 solubility. Factors of temperature and pressure were also studied. The determination of CO2 concentration in the solution was measured by gas analyzer. The result obtained from optimization revealed a maximum CO2 concentration of 19.029 mol/L in the culture media with MEA, at a pressure of 136.728 kPa, operating at 20.483 ◦C.

**Keywords:** carbon dioxide; culture media; microorganism; optimization

#### **1. Introduction**

Fossil fuels are broadly acknowledged as being the principal source of energy, and since the First Industrial Revolution the amount of carbon dioxide (CO2) in the atmosphere has risen from 280 μmol/mol to 412 μmol/mol. The resulting CO2 emissions contribute significantly to worldwide climate change [1]. Up to now, the deployment of cutting-edge low-carbon fossil-energy technologies was considered to be the ultimate solution. Preventing worldwide climate change can be achieved by taking two long-term emission objectives into account. First, CO2 emissions have had to reach their highest point, and then in the second half of the century, the goal has had to be to strive to achieve net greenhouse gas neutrality, by balancing anthropogenic emissions by the sources with the removal by sinks [2]. Hence, it is crucial to decrease such anthropogenic emissions. Second, CO2 can be captured and used as a significant feedstock to produce valuable commodities. As the world population increases, the need for energy supply rises at an exponential rate. Subsequently, to meet this demand, new and renewable energy sources are required. Along this line, treating CO2 as a feedstock to many value-added chemicals and fuels addresses both emission-control and energy supply challenges [3].

The concepts mentioned above are commonly used in carbon management from a climate change perspective. The term used is CO2 capture, utilization, and sequestration (CCUS). The carbon capture and storage (CCS) approach in reducing CO2 emissions is particularly common nowadays [4]. It refers to technologies that emphasize the selective removal of waste CO2 from a large point source, its compression into a liquified gas, and finally its transportation and sequestration to a storage site where it will not enter the atmosphere such as underground geologic formations, including depleted oil and gas reservoirs or oceans [5]. Meanwhile, carbon capture and utilization (CCU) technologies capture CO2 to be recycled for an additional application. It differs from CCS in that CCU does not permanently sequester the CO2 waste, but rather, treats it as a renewable carbon feedstock to complement the conventional petrochemical feedstocks for conversion into other substances or products with higher economic value [6].

However, due to the thermodynamically stable nature of CO2, utilizing it in chemical reactions is challenging. High energy input is required to breakdown carbon atoms in CO2 molecules, which is one of the reasons why CO2 is not extensively used in current chemical industries. Nevertheless, autotrophic microorganisms are well-known for their ability to utilize light to fix atmospheric CO2 during the process of photosynthesis. These microorganisms can capture energy in the light cycle and store it for converting adenosine diphosphate (ADP) and nicotinamide adenine dinucleotide phosphate (NADP) into adenosine triphosphate (ATP) and nicotinamide adenine dinucleotide phosphate hydrogen (NADPH) respectively. They then utilize these energy molecules during a dark cycle for transforming CO2 into valuable organic compounds [7]. Research has been done recently on altering the molecules of autotrophic cyanobacteria and algae through metabolic engineering to take advantage of their abilities to treat CO2 [8]. Microorganisms require macronutrients, micronutrients, and vitamins to grow [9]. Based on these requirements, a culture broth is required as a growth medium in a closed system, such as a bioreactor, for the production of biomass or organic compounds [10]. Therefore, increasing the CO2 solubility in a culture broth is an important step toward CO2 utilization by microorganisms.

Al-Anezi et al. studied the effect of temperature, salinity, and pressure on CO2 solubility in different aqueous solutions [11]. The relationship between these parameters on CO2 solubility was presented. Where gas solubility reduced between the temperatures of 25 ◦C and 60 ◦C, the effect was less evident at a higher temperature. Meanwhile, higher pressure (one to two bars) resulted in higher gas solubility. Additionally, the study stated that gas solubility decreased as salt increased in the solution. Yincheng et al. compared the CO2 removal efficiency of sodium hydroxide (NaOH) and aqueous ammonia [12]. The study involved the capture of CO2 in a spray column, and a fine spray of ammonia and NaOH was used. The key finding of the study was the value of mole ratios of NaOH and ammonia to CO2 suitable for the spray column, which is 4.43 and 9.68 respectively. Additionally, Martins da Rosa et al. researched the CO2 fixation of *Chlorella* using monoethanolamine (MEA) [13]. Through this research, it was found that the CO2 intake was higher for the growth of algae using a certain mass concentration of MEA; 50 mg/L and 100 mg/L for this particular strain of algae. However, the treatment of CO2 decreased if the MEA concentration used was very low or very high. Thus, the concentration of MEA used was dependent on the microorganism used.

The present paper will investigate various features that determine the concentration of CO2 dissolved in a culture solution. First, to determine the maximum CO2 concentration in the NaOH aqueous solution as a comparison, a steady rate of CO2 was supplied for a certain time. NaOH was chosen because the CO2 absorption capacity of NaOH solution is high, with a mass ratio of capture, *w*(NaOH/CO2) equal to 0.9 [14]. Second, MEA and culture media were used as the absorbent to determine the capability of both solutions in capturing CO2. Next, different kinds of culture media solutions were prepared; with the addition of either NaOH or MEA, and the absorption was carried out under the same conditions as the previous run in a batch reactor. From the experimental results, the absorption behavior is presented according to pH and time. Then optimization was run by software to determine the optimized condition for CO2 absorption.

#### **2. Materials and Methods**

#### *2.1. Carbon Dioxide (CO2) Delivery System*

A batch-typed glass (borosilicate) cylindrical reactor (Bio Gene®, Australia) with a built-in motor, the total volume capacity of5L(*D* = 140 mm; *h* = 325 mm), equipped with a pressure gauge, (RS Components, Johor Bahru, Johor, Malaysia), pH (BOQU®, Shanghai, China) and temperature probe (DPSTAR Manufacturing Sdn. Bhd., Kuala Lumpur, Malaysia) was employed for the carbon dioxide (CO2) absorption as shown in Figure 1. The reactor was connected to a pressurized gas mixture tank (Gaslink Industrial Gases Sdn. Bhd., Puchong, Selangor, Malaysia) through a flowmeter (HERO TECH®, Puchong, Selangor, Malaysia) with a valve for controlling the flow rate of the mixture. The compositions of the gas mixture were 90% CO2 and 10% Nitrogen (N2). For providing a vacuum space inside the reactor, a vacuum pump (vacuubrand®, Wertheim am Main, Baden-Württemberg, Germany) was mounted with a valve. The gas analyzer (Geotech Environmental Equipment Inc., Denver, CO, USA) was connected through one of the openings to determine the amount of CO2 in the headspace. The unwanted opening at the surface of the reactor was closed by a stopper. Additional CO2 diffuser (FunPetAqua.my, Kuala Lumpur, Malaysia) was employed to increase the CO2 absorption rate. All dimensions and components of the diffuser are described in Figure 2. All components were made of borosilicate except for the distributor head that was made of porous rubber.

**Figure 1.** Carbon dioxide absorption system.

**Figure 2.** Carbon dioxide diffuser.

#### *2.2. Time Measurement of The Maximum Carbon Dioxide (CO2) Dissolved*

An aqueous solution with the concentration of sodium hydroxide (NaOH) equal to 0.1 mol/L was prepared by dissolving sodium hydroxide pellets (Sigma-Aldrich, St. Louis, MO, USA) in 2 L of distilled water. Before absorption was conducted, the absorbent (NaOH) temperature was maintained at room temperature, 100 kPa pressure and pH = 11.0 by adding hydrochloric acid (HCl) (Sigma-Aldrich, St. Louis, MO, USA) or NaOH to adjust the pH value to a desirable number. All equipment, including tubes, fittings, and the headspace of the reactor, were sufficiently washed by a vacuum pump. After the conditions are met, the absorption was carried out by injecting the gas mixture into the absorbent via a sparger with a flow rate of 4.5 L/min controlled by a mass flow controller. The solution was mixed using a mechanical stirrer at a speed of 180 rpm for uniform reaction in an absorber. The variations of pH during the reaction were measured every 10 s. The gas mixture was bubbled into the absorbent until the pH value stops dropping. The time for the CO2 component in the mixture to dissolved in the medium was taken and a graph for pH drop against time was drawn. The same experiment was repeated while using 0.1 mol/L monoethanolamine (MEA) (Sigma-Aldrich, St. Louis, MO, USA) solution to determine the pH drop for both absorbents.

Then the experiment was repeated by replacing the absorbent with culture media. Culture media composed of 3.0 g/L dipotassium phosphate (K2HPO4), 0.2 g/L magnesium chloride (MgCl2), 0.2 g/L calcium chloride (CaCl2), and 1.0 g/L sodium chloride (NaCl) was prepared (Sigma-Aldrich, St. Louis, MO, USA). A total of 0.1 mol/L aqueous NaOH solution or 0.1 mol/L MEA solution was added to culture media to promote CO2 absorption. It is important to determine media capability as CO2 absorbent as culture media will help microbes to utilize CO2 for producing the chemical commodity. The parameter is set as the previous run, until the CO2 concentration is maximized, and then the gas mixture supply will be stopped. The graph of pH drop against time for both compositions was plotted to determine the significance of each composition. Then, all the experiments were repeated with the deployment of a CO2 diffuser at the sparger to observe the change in time for pH drop.

#### *2.3. Optimization through Surface Response Methodology*

Additionally, three factors that affect CO2 solubility (*s*), composition (*X*), pressure (*p*), and temperature (*T*) were also studied using three-factor, three-level Box-Behnken design (BBD). Each of these independent factors divided into three different levels as shown in Table 1. To describe the relationship between a set of parameters and output, the regression model was developed in BBD design and can be defined by Equation (1):

$$Y = \beta\_0 + \sum\_{i=1}^{3} \beta\_i X\_i + \sum\_{i=1}^{3} \beta\_{ii} X\_{ii}^2 + \sum\_{i=1}^{2} \sum\_{j=i+1}^{3} \beta\_{ij} X\_i X\_j \tag{1}$$

where *Y*: response; β0: constant-coefficient; β*<sup>i</sup>* and β*ii*: linear and quadratic coefficients for the terms *Xi* and *Xii*, respectively; β*ij*: coefficients which represent the interactions of *Xi* and *Xj*. Then, an analysis of variance (ANOVA) was used to determine whether the models are acceptable for analysis.


**Table 1.** Levels and ranges of independent input parameters in the Box-Behnken design.

#### *2.4. Analytical Methods*

Dissolved carbon dioxide (CO2) in culture media was determined by calculation based on readings obtained from a gas analyzer (Geotech EnvironmentalEquipment Inc., Denver, CO,USA). Readings of pH and temperature were obtained by direct measurement using a portable pH (BOQU®, Shanghai, China) and temperature probe (DPSTAR, Kuala Lumpur, Malaysia). Time was recorded using a stopwatch.

#### *2.5. Model Description and Calculation*

Dissolved carbon dioxide (CO2) in culture media can be calculated using the information given by the gas analyzer. The reading given by the analyzer consists of the composition of air in the headspace. Since the value given by the analyzer is in percentage, some calculations need to be done. The percentage of CO2 was multiplied by the volume of headspace to get the volume CO2 in the headspace in m3. The volume of headspace was constant throughout the experiment. It was set at 2.5 <sup>×</sup> <sup>10</sup>−<sup>3</sup> m3. By subtracting the total value of CO2 supplied with the one in the headspace, the value of carbon dioxide aqueous, CO2(aq) can be obtained.

#### **3. Results and Discussion**

#### *3.1. Relationship between Carbon Dioxide (CO2) Concentration and pH*

The pH of the absorbent decreased with increasing CO2 concentration. A slight difference in the lowest value of pH was observed between the absorbent used; sodium hydroxide (NaOH), culture media, and culture media with either NaOH or monoethanolamine (MEA), but the difference was not significant except for MEA which was observed to take a longer time to achieve minimum pH. A significant difference was only in the time taken to achieve the minimum value of pH. The graph of pH against time was plotted for NaOH, MEA, and culture media in Figure 3. The pH decreased with time until the 18 min mark for NaOH and the final pH was 6.91. Meanwhile for MEA and culture media were *t* = 60 min; pH = 8.52 and *t* = 8 min; pH = 6.46 respectively. Finally, culture media with NaOH and MEA were *t* = 9 min; pH = 6.45 and *t* = 21 min; pH = 7.0 respectively.

**Figure 3.** Effect of pH change for five absorbents at *p* = 100 kPa, *T* = 20 ◦C, with an initial pH of 11 with exception of culture media with an initial pH of 8.5.

Since the behavior of culture media and MEA absorbents do not follow the trend of the other three absorbents, it not suitable to be included in the optimization process (Table 1). The value of pH for culture media cannot be increased without the addition of other substances, thus, the comparison cannot be done as the initial pH value was way lower than other absorbents. For the case of MEA, the optimization process is used to find the best solution for a range of data, a large difference in data value can resulting in invalid optimization. Because of that, these two absorbents were not included in the optimization analysis. Additionally, the focus was always on culture media with the addition of NaOH and MEA.

Many factors can affect pH in the absorbent. The decrease in pH value in all absorbents was due to the increase of the concentration of dissolved inorganic carbon such as CO2(aq), HCO3 <sup>−</sup>, and CO3 2− present in CO2 that been dissolved in said absorbents [15]. Theoretically, NaOH will yield a higher CO2 solubility than culture media present because the total alkalinity of NaOH is higher due to NaOH is a strong alkaline. The mechanism of CO2 absorption in NaOH aqueous solution is summarized in the following [16]. All Na<sup>+</sup> and OH<sup>−</sup> was ionized in pure water, then aqueous CO2 reacts with OH<sup>−</sup> as expressed in the following equation:

$$\text{HCO}\_2\text{(aq)} + \text{OH}^-\text{(aq)} \rightleftharpoons \text{HCO}\_3^-\text{(aq)}\tag{2}$$

$$\rm HCO\_3^-(aq) + OH^-(aq) \rightleftharpoons H\_2O(l) + CO\_3^{2-}(aq) \tag{3}$$

Both equations are reversible reactions with a high effect on pH changes. The reaction is continuous so every CO2(aq) that was present in the medium will instantaneously be consumed. Due to high alkalinity, reaction Equation (3) was the main reaction occur in the absorbent early on resulting in increasing of CO3 <sup>2</sup>−, while OH<sup>−</sup> is rapidly consumed via both reactions. This explained the reason for the sudden change in pH at the early stage of the experiment. As CO2 aerated through the medium, OH<sup>−</sup> will keep decreasing and CO3 <sup>2</sup><sup>−</sup> keep accumulating. This phenomenon will force a backward reaction of Equation (3) which will accelerate the forward reaction of Equation (2). The pH will keep dropping in this stage. At a certain point in the experiment, pH will stop dropping and remain constant due to the reaction at equilibrium. After all the reactions are at equilibrium, the overall reaction of NaOH with CO2 can be written as Equation (4).

$$\text{NaOH(aq)} + \text{CO}\_2(\text{g}) \to \text{NaHCO}\_3(\text{aq})\tag{4}$$

Meanwhile, for the culture media, the phenomenon of CO2 dissolved can be explained by the reaction between CO2 and water (because H2O is present in media). Firstly, aqueous CO2 will react with water to form carbonic acid [17]:

$$\rm{CO\_2(aq)} + \rm{H\_2O(l)} \rightarrow \rm{H\_2CO\_3(aq)}\tag{5}$$

Then the H2CO3 can lose one or both of its H<sup>+</sup> to form:

$$\rm H\_2CO\_3(aq) \rightleftharpoons HCO\_3^-(aq) + H^+(aq) \tag{6}$$

$$\mathrm{HCO\_3^-(aq)} \rightleftharpoons \mathrm{CO\_3^{2-}(aq)} + \mathrm{H^+(aq)}\tag{7}$$

The pH drops in media are due to the released hydrogen ions. However, this equation can operate in both directions depending on the current pH level, working as its buffering system. At a higher pH, this bicarbonate system will shift to the left, and CO3 <sup>2</sup><sup>−</sup> will pick up a free hydrogen ion [18]. But in the system, CO2 was constantly added to the media, then increasing the value of dissolved CO2 causing the reaction Equations (6) and (7) forced to be carried out from left to right. This increases H2CO3, which decreases pH.

Additionally, based on Figure 2, it shows that culture media with NaOH was the fastest to reach the minimum or equilibrium pH, following by pure NaOH and media with MEA. When the pH reaches its equilibrium point, it indicates that no more CO2 is being absorbed. With this information, it is concluded that the longer time is taken to reach equilibrium, the higher the CO2 concentration in absorbent. As expected, culture media capability to absorb CO2 is weaker than NaOH. However, when MEA was added to culture media, a higher amount of CO2 was absorbed.

#### *3.2. System Performance with Deployment of Carbon Dioxide (CO2) Di*ff*user*

One of the main factors in the CO2 absorption rate is the surface area of the gas–liquid boundary [19]. In the next set of experiments, the factor was been investigated. CO2 diffuser was mounted at the sparger at the bottom of the reactor where the CO2 is aerated. This diffuser functions to turn the CO2 bubble into a more refined form. Parameters used were based on the previous experiments. The result is shown in Figure 4.

**Figure 4.** Effect of diffuser on pH change for five absorbents at *p* = 100 kPa, *T* = 20 ◦C, with an initial pH of 11 with exception of culture media with an initial pH of 8.5.

From Figure 4, the addition of the diffuser favorably affected the CO2 absorption rate by all absorbents. The final pH value were almost identical with previous run, while the time taken for pH to drop for NaOH (pH = 6.88, *t* = 15 min), MEA (pH = 8.49, *t* = 50 min), culture media (pH = 6,4, *t* = 6 min), culture media with NaOH (pH = 6.43, *t* = 7 min) and culture media with MEA (pH = 7, *t* = 17 min) were approximately decrease by 20% when using diffuser compared to the previous. By using a diffuser, CO2 is dissipated into countless small bubbles, which flows in the culture media in the form of a bubbling stream. This process is called atomization. The large number of bubbles scattered in culture media will increase the contact area of gas–liquid that will increase the absorption rate.

A simple relation between the contact surface area and mass transfer rate can be expressed in Equation (8) [20]:

$$m = \int\_{S} j\_{\rm m} \mathrm{d}S = S j\_{\rm m} \tag{8}$$

where *m*, mass transfer, *j*m, mass flow, and *S*, interfacial area, and it is shown that the relationship between mass transfer is directly proportional to the contact area. Additionally, a more detailed mathematical model of absorption rate with other parameters had been derived by Martinez I. et al. [21] using Fick's Law shown in Equation (9):

$$j\_{\mathfrak{F}} = \frac{q\_{\mathfrak{m}}}{4\pi r^2} = -D\_{\mathfrak{k}} \frac{\mathbf{d}(w\rho\_1)}{\mathbf{d}r} \tag{9}$$

where *j*g: diffusion mass flux; *q*m: mass flow rate; *D*g: gas diffusivity; *w*: mass fraction; ρ1: density of the liquid used. The assumption is as follows: symmetry is spherical, time-independent, constant liquid and gas density, and residence time of bubble much smaller than the dissolution time, the factor that affects the absorption rate can be defined by Equation (10):

$$w\_{\infty}(t\_{\rm v}) = w\_0[1 - \exp(-Kt\_{\rm v})], \text{ where } K \equiv \frac{27v\_1 D\_{\rm g} u\_{\rm g}}{2gr\_0^4} \tag{10}$$

*w*∞: mass fraction before venting; *w*: gas mass fraction; *v*: kinematic viscosity of liquid; *D*g: diffusion coefficient of CO2 gas in water; *u*g: gas injection speed; *g*: gravity acceleration; *r*0: radius of the bubble. The benefit of this simple mathematical formula is that the relations between different parameters are explicit. Gas diffusion from bubbles to the aqueous phase is measured by *w*∞, hence based on this equation, the mass of gas in a liquid is inversely proportional to the fourth power of bubble radius. This experiment only portrays the benefit of a simple diffuser. More complications, especially with made CO2 diffusers for CO2 absorption in the reactor can further increase the absorption rate of CO2.

#### *3.3. Factors A*ff*ecting Carbon Dioxide (CO2) Solubilities in Culture Media*

CO2 is one of the key factors in organic acid fermentation [22,23]. Organic acid had a wide application ranging from preservative agents for food to lab application [24]. Due to its benefit, the production of organic acid is widely studied. As mention earlier, one of the three factors that affect CO2 solubilities is the composition of media. Many studies focusing on organic acid production have increased the CO2 availability by the addition of chemicals such as magnesium carbonate (MgCO3), sodium bicarbonate (NaHCO3), or calcium carbonate (CaCO3). Through the addition of such chemicals, the CO2 solubility is greatly increased. Thus, studies on different chemicals such as sodium hydroxide (NaOH) and monoethanolamine (MEA) are also important.

The theoretical access to CO2 solubilities in culture media is very limited. The effect of organic solutes on gas solubility can be rather complex. But parameters affecting the solubilities were well-studied. Three parameters affecting CO2 solubilities are pressure, temperature, and media composition [25]. The effects of these parameters on the solubility of a gas in a pure solvent, expressed in a mathematical model were simplified in Table 2. Based on Table 2, composition (*X*), pressure (*p*), and temperature (*T*) had been identified as the three most important independent parameters affecting CO2 solubility (*s*) in absorbent and thus are chosen as the inputs for the design.


**Table 2.** Mathematical model of effects of pressure, temperature, and composition on gas solubility.

#### 3.3.1. Design of Experiment Analysis

In this study, a three-factor, three-level Box-Behnken design (BBD) was used to investigate the effects of composition (*X*), pressure (*p*), and temperature (*T*) the interactions of these factors on the CO2 solubility (*s*) in absorbent measured by its concentration. A total of fifteen experimental samples were required for the BBD including three replicated experimental runs using the processing parameters at the center points (Table 3).


**Table 3.** Actual response of CO2 concentration at experimental design points.

\* Replicated experimental runs.

The ANOVA analysis is performed as shown in Table 4 to determine the significance and adequacy of the regression models. The Model *F*-value of 135.76 implies the model is significant. There is only a 0.01% chance that an *F*-value this large could occur due to noise. *p*-values less than 0.05 indicate model terms are significant and all insignificant model terms had been reduced. Additionally, a graph in Figure 5 shows a good agreement between the actual data and the predicted values from the regression models. The Predicted *R*<sup>2</sup> of 0.9375 is in reasonable agreement with the Adjusted *R*<sup>2</sup> of 0.9897; i.e., the difference is less than 0.2. For adequate precision measures of the signal to noise ratio, a ratio greater than 4 is desirable. The ratio of 37.693 indicates an adequate signal. This model can be used to navigate the design space. This result shows that the regression model is statistically significant and adequate for the prediction and optimization of the CO2 absorption process.



*R*2: 0.9971; adjusted *R*2: 0.9897; predicted *R*2: 0.9375; Adequate Precision: 37.6933.

**Figure 5.** Comparison of actual and predicted results of CO2 absorption.

Quadratic equations were derived to describe the relationships between the parameters and CO2 solubility (*s*) as shown in Equations (11)–(13).

NaOH:

$$s = 0.001291(p)^2 + 0.019667(T)^2 - 0.298667p - 1.46100T + 53.38333 \tag{11}$$

Culture media with NaOH:

$$s = 0.001291(p)^2 + 0.019667(T)^2 - 0.301067p - 1.2235T + 44.5975 \tag{12}$$

Culture media with MEA:

$$s = 0.001291(p)^2 + 0.019667(T)^2 - 0.249667p - 1.495T + 51.4075 \tag{13}$$

3.3.2. Individual Effect of Experiment Parameters on Carbon Dioxide (CO2) Absorption

The significance of each parameter on CO2 solubility can be illustrated by the perturbation plot in Figure 6, where steep slope and curvature were obtained for both pressure and temperature indicates that CO2 solubility is sensitive to the parameters. The maximum value of CO2 dissolved achieved with the addition of NaOH for culture media was lower than what been achieved for MEA. This is due to the MEA's rapid reaction with CO2 in low partial pressure [27].

**Figure 6.** Perturbation plot comparing the response of CO2 solubility to changes in temperature and pressure in (**i**) pure NaOH, (**ii**) culture media with NaOH, and (**iii**) culture media with monoethanolamine (MEA).

Figure 6 shows the relationship between pressure and final CO2 concentration which is directly proportional which was described quantitatively by Henry's law shown in Table 2; pressure. External pressure affects the concentration of gas molecules in space. When the partial pressure of the gas above the solution increases, it forces the gas molecules to solute in solution to maintain dynamic equilibrium [28]. Additionally, high temperature demotes CO2 absorption as opposed to high pressure. Also, the effect of temperature is more significant than the pressure (highest *F*-value). A slight increase in temperature will greatly reduce CO2 absorption. While to achieve more significant change by pressure, extremely high pressure needs to be applied [29]. The gas dissolves in liquid because of the interactions between its molecules and absorbent. This interaction will release heat when these new attractive interactions form in an exothermic process. Thus, additional heat will produce thermal energy that overwhelms the attractive forces between the gas and the absorbent molecules resulting in less CO2 dissolved in solution [28].

#### 3.3.3. Collective Effect of Experiment Parameters on Carbon Dioxide (CO2) Absorption

The *F*-value determine the relative importance of a parameter. From Table 4, the temperature has the highest *F*-value, which is 446.76, showing its importance in CO2 absorption in this case. Figures 7–9 are three-dimensional response surface and project contour for NaOH, culture media with NaOH, and culture media with MEA, respectively, which show the different experimental parameters and their effects on CO2 concentration in the absorbent. All graphs derived had curved rather than straight lines, indicating the strong interaction between parameters and the output.

**Figure 7.** Effect of pressure, temperature and their interaction on CO2 concentration in NaOH (**i**) three-dimensional surface plot; (**ii**) projected contour plot.

**Figure 8.** Effect of pressure, temperature and their interaction on CO2 concentration in culture media with NaOH (**i**) three-dimensional surface plot; (**ii**) projected contour plot.

**Figure 9.** Effect of pressure, temperature and their interaction on CO2 concentration in culture media with MEA (**i**) three-dimensional surface plot; (**ii**) projected contour plot.

#### 3.3.4. Process Optimization

It was observed that carbon dioxide (CO2) solubility (*s*) reaches the maximum values under low temperature operating in high pressure. However, there was a certain point in these parameters where a further change in their value will not affect the final CO2 solubility (*s*). Therefore, a balance value needs to be established to achieve the maximum result while optimizing the parameters. The purpose of this step is to observe the combination of the independent variable (i.e., absorbent, temperature, and pressure) to get the maximum CO2 solubility (*s*) simultaneously. The overall performance of the CO2 absorption strongly depends on a wide range of experimental conditions. It is crucial to optimize the CO2 absorption process with multiple inputs and multiple responses. Thus, a series of optimizing results proposed by response surface methodology (RSM) with multiple inputs and multiple responses are listed in Table 5. The optimal CO2 solubility (*s*) can be obtained in the following conditions: culture media with MEA as absorbent; 20.483 ◦C of temperature; 136.728 kPa of pressure. The optimal CO2 solubility (*s*) obtained at this condition was 19.029 mol/L. Additionally, a favorable value for each parameter can be chosen from the RSM. For example, the desired temperature is ≈ 22 ◦C, pressure must be around 140 kPa to obtain the optimal CO2 solubility (*s*). Though from the optimization process, absorbents other than culture media with MEA was found to be undesirable.


**Table 5.** Process optimization for CO2 absorption by RSM.

#### **4. Conclusions**

The present work has investigated the features of sodium hydroxide (NaOH) aqueous solution and culture media to capture carbon dioxide (CO2) at different compositions, pressure, and temperature. It was observed that the CO2 solubility increases using monoethanolamine (MEA) compared to NaOH. CO2 absorption was also favorable at high pressure and low temperature. Improvement of the absorption rate can be achieved by deploying a CO2 diffuser. Using optimization software, the most optimized condition for the CO2 absorption process was by using culture media added with MEA, at a pressure of 136.728 kPa operating at 20.483 ◦C. Besides, capturing CO2 by using culture media will lead to the production of chemical commodities (such as succinic acid, formic acid, and acetic acid) by microbes that can be useful for industrial usage. This work serves as a base for further research on CO2 absorption by culture media. Further studies such as the feasibility of this method by using gas mixture instead of pure CO2 are necessary to demonstrate a more flexible process. The ultimate aim of this work is to produce value-added commodities utilizing CO2 as the main carbon source with the help of suitable microbes.

**Author Contributions:** Conceptualization—S.N. and A.I.A.; writing, original draft preparation—S.N. and A.I.A.; writing, review, and editing—M.Y.O. and S.N.; writing, proofreading—A.I.A. and P.L.S.; funding acquisition—S.N. All authors have read and agreed to the published version of the manuscript.

**Funding:** The authors (SN and AIA) would like to thank the Ministry of Education (MOE) Malaysia for the financial support through the Fundamental Research Grant Scheme (MOHE Project Ref. No.: FRGS/1/2018/STG01/UNITEN/01/1) and BOLD 2025 (10436494/B/2019095). AIA also wishes to express his gratitude to UNITEN for providing the UNITEN Postgraduate Excellent Scholarship 2019.

**Acknowledgments:** The authors would like to express their special thanks to Universiti Tenaga Nasional for providing facilities and equipment to ensure the accomplishment of this project.

**Conflicts of Interest:** The authors declare that they have no competing interests.

#### **References**


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### *Article* **E**ff**ects of Alginate and Chitosan on Activated Carbon as Immobilisation Beads in Biohydrogen Production**

**Nur Farahana Dzul Rashidi 1, Nur Syakina Jamali 1,\*, Siti Syazwani Mahamad 1, Mohamad Faizal Ibrahim 2, Norhafizah Abdullah 1, Siti Fatimah Ismail <sup>1</sup> and Shamsul Izhar Siajam <sup>1</sup>**


Received: 28 July 2020; Accepted: 14 September 2020; Published: 6 October 2020

**Abstract:** In this study, the effects of alginate and chitosan as entrapped materials in the biofilm formation of microbial attachment on activated carbon was determined for biohydrogen production. Five different batch fermentations, consisting of mixed concentration alginate (Alg), were carried out in a bioreactor at temperature of 60 ◦C and pH 6.0, using granular activated carbon (GAC) as a primer for cell attachment and colonisation. It was found that the highest hydrogen production rate (HPR) of the GAC–Alg beads was 2.47 ± 0.47 mmol H2/l.h, and the H2 yield of 2.09 ± 0.22 mol H2/mol sugar was obtained at the ratio of 2 g/L of Alg concentration. Next, the effect of chitosan (C) as an external polymer layer of the GAC–Alg beads was investigated as an alternative approach to protecting the microbial population in the biofilm in a robust environment. The formation of GAC with Alg and chitosan (GAC–AlgC) beads gave the highest HPR of 0.93 ± 0.05 mmol H2/l.h, and H2 yield of 1.11 ± 0.35 mol H2/mol sugar was found at 2 g/L of C concentration. Hydrogen production using GAC-attached biofilm seems promising to achieve consistent HPRs at higher temperatures, using Alg as immobilised bead material, which has indicated a positive response in promoting the growth of hydrogen-producing bacteria and providing excellent conditions for microorganisms to grow and colonise high bacterial loads in a bioreactor.

**Keywords:** biohydrogen production; immobilised cells; entrapment; alginate; chitosan; activated carbon

#### **1. Introduction**

Biohydrogen is a popular energy carrier as its production promises clean energy that only generates water upon combustion, with higher energy content per unit weight (122 kJ/g) than any other fuel [1]. Biological methods have been studied to ensure the hydrogen production is safer and more economical than thermochemical methods. Dark fermentation is being recognised as an excellent biological method of hydrogen production because of its ability to perform without light energy and oxygen source [2]. Fermentation is the process of using specific microorganisms to convert organic substrates into hydrogen, carbon dioxide, and other solutes such as acetate, butanol, and ethanol [3].

The production of biohydrogen using a suspended cultivation system through high-temperature operation (50–60 ◦C) has gained attention due to its higher yield and hydrogen productivity (HPR) capability. This operation is preferable in pathogenic destruction as it can restrain the growth of hydrogen consumers such as homoacetogens and methanogens [4,5]. However, the lower microbial cell density at this temperature is a disadvantage for the fermentation. Difficulties in the retention

of biomass in the suspended system and cell washout are regularly experienced inside the reactor, which usually happens during the short hydraulic retention time (HRT) [6,7]. As a consequence, attached cell immobilisation is an approach to maximising and maintaining biomass, such that it can work at a higher rate of dilution without biomass washout from the reactor. Immobilisation technology has been developed to increase hydrogen production by providing a favourable environment of support for microbial cells during fermentation. Hence, the selection of the supporting material is imperative because it affects the overall performance of biohydrogen production.

Alginate (Alg) is an excellent support material, making it a practical choice in immobilisation. It has been reported that biohydrogen production increases three-fold when using alginate beads supplemented with aluminum oxide and titanium oxide [8,9]. Meanwhile, Wuet al. [10] said that biohydrogen production is two-times greater when alginate beads are enhanced with activated carbon. Nonetheless, even though alginate beads have been widely used in immobilisation, they are reported to still suffer from certain limitations like weak mechanical strength and reduced porosity [11,12]. Therefore, several approaches have been studied to improve the permeability and mechanical stability of alginate matrices, such as incorporating other materials like cellulose, metal, and carbon sources.

A previous study reported that the entrapment between chitosan and alginate forms a strong ionic interaction between carboxyl groups of alginate and amino groups of chitosan, thus resulting in an improvement in the mechanical properties of the matrix support [13,14]. In other studies, the formation of high, crosslinked, porous beads, with better mechanical and chemical stability of the support matrix in the buffered medium, is produced from the ionotropic gelation of chitosan, leading to low rates of cell leakage even at higher cell loading [15]. It was also reported that the effectiveness of chitosan coating enables the physical isolation of bacteria from the outer environment and reduces cell detachment during fermentation, besides improving the mechanical strength of alginate bead carriers during storage [16–18].

The entrapment technique is widely used, which can be done in a simple procedure. This technique involves an entrapment process in which the enzyme is crosslinked within polymeric materials such as calcium alginate, polyacrylamide (PAM) gel, and agar [19]. The research reported that the stability of immobilised cells can be enhanced via the fusion of microbial cells into a rigid network of the polymer due to the mechanical firmness and good porosity of the carrier, thus providing the right anaerobic conditions to microorganisms during hydrogen production [20].

In this study, the effect of alginate and chitosan was investigated as one of the potential entrapped-material approaches in cell immobilisation and cell encapsulation. In the first part of this research, the ability of microbial culture to attach and maintain itself on a granular activated carbon (GAC) surface as their support material was performed on mixtures of glucose and xylose as the carbon source. GAC has a high surface area, low toxicity, and excellent mechanical properties, which are ideal for fermentation at high temperatures. Moreover, its characteristic of having highly porous structure helps to preserve cell viability, which serves an excellent purpose in the field of microbial colonisation where fermentative bacteria can expand freely on the surface of the supporting material and form a biofilm [21]. Efforts have been made to improve the high cell density that facilitates the good production of hydrogen. Furthermore, entrapment is part of immobilisation methods that have been used to improve the productivity of enzyme or microbial cells. This study investigated the variations of GAC–alginate (GAC–Alg)- and GAC–alginate–chitosan (GAC–AlgC)-immobilised beads, which acts as a support carrier during biofilm development in batch fermentation of biohydrogen production.

#### **2. Materials and Methods**

#### *2.1. Microorganism, GAC Carrier, Alginate, and Chitosan Carriers*

The microorganism source was collected from a sludge pit of palm oil mill effluent (POME) that was located at Sime Darby Plantation, Selangor, Malaysia. The sludge containing mixed culture underwent a heat-treated process at 80 to 90 ◦C for 60 min to prevent the development of the methanogenic population prior to use. The GAC carrier originated from shells of coconuts that were supplied by KI Carbon Solutions Sdn Bhd. GAC was sieved to attain 2–3 mm of particle size. Sodium alginate powder and chitosan flakes originated from crab shells were supplied by BT Science Sdn Bhd. Sodium alginate powder was dissolved into 1 L of distilled water and stirred using a hot plate magnetic stirrer for 30 min to attain homogeneity prior to use.

#### *2.2. Biofilm Formation on Activated Carbon*

Biofilm was primarily developed on the surface of GAC using the surface attachment method as one of the immobilisation approaches. A similar ratio of GAC to sludge, 10:10 (*w*/*v* g/L), was acclimatised in the synthetic medium inside a 1 L modified Schott bottle using the sequencing batch operation mode of 2 days HRT. The biofilm was continuously developed until biogas production was consistently obtained. The synthetic medium was used in sequencing batch fermentation. The medium contained (per liter of deionised water): KH2PO4 0.75 g L<sup>−</sup>1, NH4Cl 1 g L−1, K2HPO4.3H2O 1.5 g L−1, NaCl 2gL<sup>−</sup>1, NaHCO3 2.6 g L<sup>−</sup>1, MgCl2.6H2O 0.5 g L−1, CaCl2.2H2O 0.05 g L−1, yeast extract 2 g L−1, xylose 10 g L<sup>−</sup>1, and glucose 10 g L<sup>−</sup>1. The fermentation system was cultivated for 48 h in a water bath shaker at 60 ◦C and 120 rpm, with the pH of the culture medium adjusted to pH 6.0 [21,22].

The gas produced was monitored using the water displacement method. The measuring cylinder was put invertedly in the hydrochloric acid solution (with pH 2) to avoid the gases from being released into the environment. The volume of biogas was recorded in every cycle and collected once the stationary phase was achieved. The experimental setup of this study is shown in Figure 1.

**Figure 1.** Experimental setup for cell acclimatisation

#### *2.3. Development of GAC—Attached Biofilm Entrapped in Alginate Beads (GAC–Alg)*

The different concentrations of alginate were prepared by dissolving 0.5, 1, 2, 3, and 4 g of sodium alginate powder into 1 L of distilled water, as shown in Table 1. About 40 g of GAC-attached biofilm were put into the alginate solution and mildly stirred until well mixed. The mixed granules were then dropped into a 2% (*w*/*v*) solution of 100 mL of calcium chloride (CaCl2) in a separate beaker to form and harden the beads and left for 30 min. The hardened beads, obtained with a diameter range of approximately 4–5 mm, were filtered and rinsed with sterile water before use.


**Table 1.** Samples of granular activated carbon–alginate (GAC–Alg)- and granular activated carbon– alginate–chitosan (GAC–AlgC)-immobilised beads *v*/*w***.**

#### *2.4. Entrapment of GAC–Alg Beads with Chitosan (GAC–AlgC)*

The previous method of GAC-attached biofilm with alginate was repeated. The different concentrations of chitosan were prepared by dissolving 0.5, 1, 2, 3, and 4 g of chitosan flakes into 5% (*v*/*v*) of acetic acid (5 mL) in a separated beaker, as shown in Table 1. The beads formed were added into chitosan solution until they were well immersed. The beads were then sunk into 40 g of NaOH for 30 min to make it hardened and fully coated before being filtered and rinsed with sterile water and used in fermentation [23,24].

#### *2.5. Batch Fermentation of Biohydrogen Production*

The fermentation process was carried out in a 250 mL Schott Duran bottle, as shown in Figure 1 Nitrogen gas was pumped into the bottle for 2 min before fermentation to eliminate the oxygen inside the bottle. The fermentation process was carried out for 12 h, with an initial medium of pH 6.0, as well as temperature and shaking speed at 60 ◦C and 120 rpm, respectively. The process was repeated periodically for two batches with different types of substrates [25,26]. The gas samples generated during the fermentation were collected when the biogas amount was consistently achieved.

#### *2.6. Analysis of Gaseous, Hydrogen Yield, and Productivity*

The hydrogen yield (HY) was determined based on the amount of hydrogen produced over the amount of sugar consumed. Hydrogen yield was represented as hydrogen moles per mole of sugar consumed. The percentage of biogas composition was examined using gas chromatography (GC) (Model HP6890N, Agilent Technology, USA) consisting of two detectors: a thermal conductivity detector (TCD) andflame-ionization detector (FID). The internal diameter and film thickness of the column was 0.53 mm and 0.5 mL, respectively. The oven temperature was set at 75 ◦C, and the carrier gas flow rate (argon) was 6 mL/min. Then, 0.5-mL samples of gas were taken using a 1-mL gas-tight syringe, injected into the GC immediately. The TCD was calibrated with standard gas (Air Product, Malaysia) mixtures, consisting of H2, CH4, CO, and CO2 in nitrogen, at periodic intervals.

Modified Gompertz was presented to correlate the cumulative hydrogen gas production using the Solver add-in in Excel. Theoretically, the Gompertz equation was modified [26]

$$H\_t = \; H\_{m.c} \exp\left\{-\exp\left[\frac{R\_{m.c}}{H\_m}(\lambda - t) + 1\right]\right\}.\tag{1}$$

where *Ht* is the cumulative hydrogen production (mL), *Hm* is the maximum hydrogen production (mL), *Rm* is the maximum hydrogen production rate (mL.h<sup>−</sup>1), *<sup>e</sup>* is the Euler number (*<sup>e</sup>* <sup>=</sup> 2.73), <sup>λ</sup> is the lag phase time (h), and *t* is the incubation time (h).

#### *2.7. Analysis of Volatile Fatty Acid and Sugar*

HPLC analysis was used to determine the number of monosaccharides mainly found as xylose and glucose, and also the amount of volatile fatty acids (TVFAs) that was present in a sample. The liquid samples were filtered into vials via a 0.22-μm syringe. The soluble microbial product (SMP) and monomeric sugar concentrations were quantified by HPLC analysis fitted with a refractive index detector (RID) with a column (Phenomenex, RPM Pb2+). The mobile phase used was 5 mM water at a constant flow rate of 0.6 mL/min at room temperature. The column temperature was maintained at 80 ◦C, and the HPLC sample injection volume was 20 μL. The intended compounds were identified by conducting standard curves of different concentrations of SMPs and sugar concentrations.

#### *2.8. Scanning Electron Microscope (SEM)*

The formation of cell attachments on the immobilisation beads was observed by using scanning electron microscopy (SEM) [27]. The appearance of beads was seen before and after the fermentation process. The size of both types of beads was measured as 4–6 mm per bead. The physical stability of the beads was observed by putting the bead samples separately into test tubes with a medium of synthetic solution at pH 6.0 and keeping them in the same water bath shaker for fermentation. The state of the physical changes of the beads was recorded after they began to degrade. For further analysis, the beads were taken right after the fermentation process and left at −20 ◦C before the morphology test. The experiment was conducted using a scanning electron microscope (SEM; model Q250, Thermo Scientific, Waltham, MA, USA). The beads were cut into half with a knife to inspect the structure inside. The gel beads were then mounted on metal stubs, and the inside layer underwent sputter-coating with gold for 6 min. Then, the surfaces were examined and captured.

#### **3. Results and Discussion**

#### *3.1. Microbial Cells Self-Attached to GAC for Hydrogen Production*

The microbes were cultivated in a synthetic medium containing glucose and xylose mixtures as the sole carbon and energy source until biogas production was consistently achieved. The ability of the cells to bind themselves (self-attach) to the GAC surface had been thoroughly evaluated. Biogas production (mL) was plotted over fermentation time (day), as shown in Figure 2. It can be seen that the biogas fluctuated over the fermentation period and started to be consistently produced at Day 30, towards the end of 40 days of fermentation, with cumulative biogas production being 2115.75 ± 413.03 mL and 2274.75 ± 411.83 mL for 10 and 20 g/L sugar loading, respectively. The results of the production of biogas via dark fermentation immobilised with GAC are shown in Table 2.

**Figure 2.** Biogas production (mL) of 10 g/L and 20 g/L sugar loading over fermentation time (day) with 2 days HRT in sequencing mode reactor.


**Table 2.** Hydrogen productivity and H2 yield obtained from each of the different sugar loadings.

The process continued until the biogas was stable and ready for gaseous analysis by using GC. The average hydrogen production rate was recorded as 3.71 ± 0.09 mmol H2/l.d at 10 g/L sugar used and 3.92 ± 0.21 mmol H2/l.d at 20 g/L sugar used. This indicates that 20 g/L is the optimal amount of sugar to be used for immobilisation beads and, thus, as the optimum substrate for future experiments. In parallel, our previous work also suggested that 20 g/L sugar was the optimal amount of sugar to use [21]. From the data obtained, it was found that the granular activated carbon could provide a suitable matrix to become a primer for cell attachment and colonisation before entrapment for hydrogen production. This is due to the mechanical stability of the biofilms formed on the activated carbon, which have a high propensity in binding capacity, providing a nutrient-rich environment, and thus promoting microbial adhesion [28,29]. The attachment-formed biofilms also help to sustain cell viability and prevent cell washout from the reactor, thus increase cell density [30].

#### *3.2. Biohydrogen Production of Immobilised Beads GAC–Alg*

The different concentrations of alginate (Alg) ratios were added to GAC to determine an optimum amount of alginate for biohydrogen production. The results of hydrogen gas produced in each run were plotted against time. From Section 3.1, the optimum result was obtained when 20 g/L amount of sugar was used as a substrate in this experiment, which was subjected to 20% *w*/*v* of GAC–Alg as immobilised beads in 200 mL working volume.

Hydrogen production using the entrapment technique as immobilised beads was evaluated. Figure 3 shows the comparison of biogas trend production for GAC–Alg beads during the acclimatisation period using a synthetic medium as a substrate. Hydrogen production started to increase at 4 h of fermentation for five different concentrations of alginate, dominantly by GAC–Alg beads at C with a ratio of 1:2.

**Figure 3.** Hydrogen production (mL) at different concentrations of alginate in 200 mL of a 250-mL modified bioreactor in batch fermentation.

The results of HPR (mmol H2/l.h) and hydrogen yield (mol H2/mol sugar) were plotted against different concentrations of GAC–Alg, as shown in Figure 4. It can be seen that the highest hydrogen production was found for C at the GAC–Alg ratio of 1:2, with HPR of 2.47 ± 0.47 mmol H2/l.h. The highest hydrogen yield was 2.09 ± 0.22 mol H2/mol total sugar run at C. The cell density (in VSS) of the GAC–Alg was produced at the highest value of 1.65 g/L, as compared to the density of the lowest concentration of GAC–Alg at Run A, which was only 0.48 g/L.

**Figure 4.** Hydrogen productivity rate (mmol H2/l.h) and hydrogen yield (mol H2/mol sugar) in the different concentrations of alginate for hydrogen production.

The trends of HPR and hydrogen yield obtained in Table 3 were comparable to the other runs that contained GAC–Alg at different ratios of B, D, E, and F. It was slightly different in terms of hydrogen yield under different concentrations of alginate, comparing the higher results of Runs B and C to Run D. It can be determined that the irregular space and porous structure present on the carrier's surface provided the microbes with ample space to develop well, in agreement with the results obtained by [31,32]. However, the trends decreased with the increment of added alginate in Runs E and F. It shows that when surrounded by a large number of support carriers, the microbial population had some limitations to grow. Increasing the concentration of alginate did not improve the beads' robustness, and the production of hydrogen gas was slower because a higher amount of alginate also acted as a barrier to the substrate and products [33].


**Table 3.** Hydrogen productivity obtained from different concentrations of alginate in batch fermentation.

The optimum ratio of GAC to alginate of 1:2 remarked the occupation of the optimal porous space of GAC–Alg immobilised beads by the microbes to promote stable biological activity for biohydrogen production. Hence, the study revealed that a combination of GAC and alginate as immobilised beads gave a positive response in bacterial immobilisation, especially in promoting the growth of hydrogen-producing bacteria during the fermentation [34]. The positive performance of the GAC–Alg beads was due to the presence of granule activated carbon inside, which acted as a support for the alginate carrier and maintained the stability of beads [21].

#### *3.3. Bacteria Immobilisation in GAC–Alg Entrapped with Chitosan on Hydrogen Production*

The development of entrapped GAC–Alg in chitosan was studied to investigate the adherence of GAC–Alg beads with regards to the mucoadhesion behaviour of chitosan. As reported by Szyma ´nska and Winnicka [35], chitosan possesses good mucoadhesion behaviour resulting from the cationic properties, existence of amino groups, and free hydroxyl, which allow the polymers to interact with each other by electrostatic and hydrogen bonding. The capability of chitosan to trap the GAC–Alg beads had been thoroughly evaluated. Different concentrations of chitosan subjected to 20% *w*/*v* of GAC–AlgC as immobilisation beads in 200 mL working volume (*w*/*v*) were used. The results of the hydrogen production (mL H2) were plotted over fermentation (hr), as shown in Figure 5. It showed the comparison of hydrogen production trends for different concentrations of chitosan g/L used during the acclimatisation period, using a synthetic medium as a substrate. Hydrogen production started to increase at 4 h of fermentation and dominantly during Run C, which had a ratio of chitosan of 1:2. The consistency of hydrogen against chitosan concentration g/L was consistent after 40 h of operation.

**Figure 5.** Hydrogen production (mL) at different concentrations of chitosan in 200 mL of a 250 mL modified bioreactor in batch fermentation.

The results of HPR (mmol H2/l.h) and hydrogen yield (mol H2/mol sugars consumed) were plotted over different concentrations of chitosan (g/L), as shown in Figure 6. The entrapment of GAC–Alg beads with varying concentrations of chitosan was measured from the evolved gas during the acclimatisation process. The results were analysed and presented in Table 4, which shows that the chitosan concentration at C with 2 g/L reached the highest level for both HPR (0.93 ± 0.05 mmol H2/l.h) and H2 yield(1.11 ± 0.35 mol H2/mol sugar consumed) with 86.63 H2 %. Meanwhile, at a lower concentration than C, which is concentration atB reached the second highest HPR of 0.85 ± 0.08 mmol H2/l.h and H2 yield of 0.97 ± 0.21 mol H2/mol total sugar with 84.79 H2 %. The beads of Run D, with 3 g/L, followed as the thirdighest HPR of 0.74 ± 0.15 mmol H2/l.h and H2 yield of 0.88 ± 0.12 mol H2/mol total sugar with 64.54 H2%. HPR of ratios A 0.5 g (as the lowest concentration) and E (as the highest concentration, with 4 g) was proportionate between those two and was recognised as causing lower hydrogen production than Concentrations B, C, and D after 52 h of operation, individually at 0.58 ± 0.20 and 52.94, and 0.70 ± 0.20 and 76.43 (mmol H2/l.h; H2%).

**Figure 6.** Hydrogen productivity rate (mmol H2/l.h) and hydrogen yield (mol H2/mol sugar) in the different concentrations of chitosan.


**Table 4.** Hydrogen productivity obtained from different concentrations of chitosan in batch fermentation.

The comparison of these findings revealed that the immobilised beads of Run C reached the highest hydrogen production and they were examined as the optimum concentration for microbial support matrix in immobilisation bead development. The work by Damayanti et. al [36] reported that among a variety of chitosan applications, chitosan in encapsulation technology is widely applied whether as a second-layer coating or in combination with other polymers. It was also reported that chitosan could improve the stability of the capsules. In other studies reported by Žuža et al. [16], it was claimed that the mechanical confidence of alginate beads increased up to seven days when coated with chitosan, which significantly contributes to the preservation of carrier strength during fermentation. The formation of the shape of cell-immobilised GAC, cell-immobilised GAC-Alg and GAC-AlgC were presented in Figure 7. Figure 7a image of cell-immobilised into GAC, (b) the shape of cell-immobilised GAC with alginate and (c) the shape of cell- immobilised GAC-Alginate beads with chitosan. It can be seen both GAC-Alg and GAC-Alg coated with chitosan were not much spherical. The GAC covered with alginate were transparent, while the beads coated with chitosan have slightly cloudy of physical appearance. Generally, the differences in the shape of beads were caused by the gravity and surface tension imbalance when the beads dropped from the syringe. The beads shape formation also were affected by the viscosity of alginate and chitosan, and distance of dropper to gel solution [34]. This is a new method introduced to improve the stability of biofilm formation and adsorption capacity as well as enhanced the mechanical strength of the carrier, thus enhanced the hydrogen yield production. Table 5 summarizes the comparison of a similar study on the efficiency of different types of immobilisation beads on hydrogen production. Table 5 shows that the carbon source and fermentation process were different to produce hydrogen gas. In the present work, the highest hydrogen yield was 2.09 mol H2 /mol sugar obtained from GAC-Alg immobilised beads. It should be

noted that the type of fermentation process and carbon source can always influence the results [36]. Likewise, the selection of the materials to be used as immobilising carriers also playing an important role in biohydrogen production, in term of high resistance towards temperature, mechanical strength as well as the possibility to recycle the immobilised carriers. Hence, in this research, attached-biofilm of hydrogen-producing bacteria on GAC from adsorption approach are stabilised using alginate entrapped with chitosan to form stable cells immobilised beads as a novel approach.

**Figure 7.** (**a**) Cell-immobilised GAC; (**b**) cell-immobilised GAC–Alg; (**c**) cell-immobilised GAC–AlgC.



Ac = activated carbon; GAC = granular activated carbon; CA = calcium alginate; C = chitosan.

#### *3.4. E*ff*ect of Alginate and Chitosan Concentration on Volatile Fatty Acid Production*

Hydrogen production performance is usually monitored collectively with the formation of acetic acid (HAc) to butyric acid (HBu) and total volatile fatty acids (TVFAs). In this anaerobic hydrogen production, the concentration of TVFAs and their relative proportions were effectively used as indicators. The plot between HPR (mmol H2/l.h) and volatile fatty acid (VFA) concentration (mg/L) for GAC-Algand GAC-AlgC-immobilised beads in batch fermentation is presented in Figure 8a for GAC-Alg and Figure 8b for GAC-AlgC. The summary of total volatile fatty acids at various concentrations of GAC-Alg and GAC-AlgC shown in Table 6

(**a**)

(**b**)

**Figure 8.** Hydrogen production rate (mmol/l.h) and VFA concentration (HAc-acetate acid and HBu-butyrate acid) in batch fermentation of (**a**) GAC-Alg and (**b**) GAC-lgC beads.



TVFAs (total volatile fatty acids) = HAc + HBu.

Glycolysis is the gateway for the metabolic process of cells that convert glucose into pyruvate as an intermediate metabolite. Pyruvate reacts to acidogenesis and generates VFAs, including butyric acid, acetic acid, and also propionic acid under anaerobic conditions [34]. Theoretically, the maximum amount of H2 yields when all glucose has been converted to HAc is 4 mol H2 per mole of glucose in Equation (2), while HBu is 2 mol H2 per glucose in Equation (3) [44].

$$\rm C\_6H\_{12}O\_6 + 2H\_2O \to 2CH\_3COOH + 4H\_2 + 2CO\_2 \tag{2}$$

$$\rm C\_6H\_{12}O\_6 + 2H\_2O \to CH\_2CH\_2CH\_2COOH + 2H\_2 + 2CO\_2 \tag{3}$$

It was found that HBu and HAc were the primary volatile fatty acids that were produced, while propionate (HPr) contributed to negligible amounts. The result of GAC–Alg dominated, with 23.99 ± 3.60 mM of HBu and 19.96 ± 2.61 mM of HAc found at Run C. For GAC–AlgC, the result was dominated by the ratio of 1:2, with 14.45 ± 1.76 mM of HBu and 12.41 ± 2.49 mM of HAc. Hydrogen production was slightly increased as the concentration of alginate increased from Runs A to C as Hbu and HAc increased. These results are close to similar metabolic pathways of hydrogen production found by [45,46], who stated that when the composition of HAc and HBu increases, the hydrogen production efficiency should also increase.

Based on these findings, it was found that HBu and HAc were eminently affected by hydrogen production in Equations (2) and (3). The highest range found in butyrate acid proposed that the biohydrogen production from a mixture of xylose and glucose was of a butyrate type in Equation (3). Hence, the investigation suggests that butyrate is significantly affected by glucose and xylose consumption rather than acetate, as reported by [45].

#### *3.5. SEM of Immobilized Beads*

Furthermore, this study observed the microbial cell culture on carriers using a scanning electron microscope (SEM). GAC as a primer for cell attachment and colonisation was observed before and after acclimatisation. The image of the micropores of clean GAC is shown in Figure 9a, while the image of Figure 9b shows the microbial cells that have successfully attached to the GAC surface. Both of the images were captured using a field emission scanning electron microscope (FESEM) at 10.00 k magnification [27]. The porosity of GAC was provided with a pleasant environment and conditions for cells to adhere themselves firmly inside the pores, thus helping the cells to grow and form the population. This overcomes the problem of self-detachment of hydrogen-producing cells during repeated batch fermentation in the culture medium [4].

**Figure 9.** Images of (**a**) clean GAC; (**b**) GAC-attached biofilm; (**c**) immobilised cells on GAC–Alg at (**i**) 3.00 k magnification and (**ii**) 10.00 k magnification; (**d**) immobilised cells on GAC–AlgC at (**i**) 3.00 k magnification and (**ii**) 10.00 k magnification.

A significant number of microbial cells were observed to have successfully immobilised into the alginate surface, as shown in Figure 9c(i), and agglomerated each other (Figure 9c(ii)). These features indicate that the alginate does not provide toxic and non-nutritive environments towards the microbial cells, but gives them a suitable place to grow dominantly, besides protecting the cells inside the beads. A part of the entrapment of cells into alginate, the image of the predominantly rod-shaped microbial species, was captured after the fermentation process, which can be clearly seen in Figure 9d(ii). The rod-shaped bacterial cells appeared to be the dominant consortium on the GAC based on their morphologic properties. This study is in agreement with a previous study by Jamali et al. [21]. It has

been identified that the dominant species of anaerobic hydrogen producers is *Thermoanaerobacterium thermosaccharolyticum*, which is stated to have spores and is rod-shaped and Gram-positive [21,28]. A similar source of inoculum was used in this research as this species has been recognised as having a high propensity to be cultivated at an optimal temperature of 50–60 ◦C and a pH of approximately 5.5 to 6.5. This bacterium played a significant role in the production of butyric acid, acetic acid, and hydrogen. This research is supported by [21], where Thermoanaerobacterium thermosaccharolyticum has been documented to be effective hydrogen producers of xylose and glucose, with butyrate and acetate as the main byproducts of fermentation in a synthetic medium. Thermoanaerobacterium thermosaccharolyticum has also been documented as capable of fermenting a broad variety of carbohydrates and complex sugars that are present in almost all wastewater [47]. The accumulation of microbial cells around the GAC–AlgC carrier surface membrane, as illustrated in Figure 9d(ii), explains the right conditions of the carrier, allowing the retention of the biological activity of the encapsulated cells. There was a significantly high mortality of cells when using chitosan as an external encapsulation agent. The hydrophobicity behaviour of chitosan was favoured by the undesired protein adsorption and denaturation process. Moreover, the diffusion issues also affected the molecular traffic of substrates and products of the microbial enzymatic process between the outsides and the insides of the carrier [48,49]. Generally, high microbial loadings hosted within the carrier showed that the cells are protected from microbial attack and physical or mechanical damage. The high porosity of the microbial matrix support will provide the right places for cells to grow and immobilise. The suitable chemical nature of carriers also can help the cells to extend as well as their protein can be easily accommodated within the channel. Nevertheless, the unavoidably fragile and prone-to-grinding environment of the carrier needs to be enhanced with optimal entrapment agents.

#### **4. Conclusions**

This work has successfully developed the entrapment of immobilised GAC with alginate and chitosan in the batch fermentation system from xylose and glucose fermentation. It was found that concentrations of alginate- and chitosan-immobilised beads at 2 g/L presented the highest amount of hydrogen productivity. The results showed that the immobilised beads maintained their stability in hydrogen production after 40 h; a consistent HPR of 2.47 ± 0.47 mmol H2/l.h and H2 yield of 2.09 ± 0.22 mol H2/mol total sugar was found with GAC–Alg beads. The consistent HPR obtained with GAC–AlgC beads was 0.93 ± 0.05 mmol H2/l.h, along with H2 yield of 0.88 ± 0.12 mol H2/mol total sugar. In accordance with all of the significant results, it is emphasised that the acclimatisation of GAC–Alg and GAC–AlgC beads as support carriers ensures the continuity of HPR and enhances cultural density in the handling of synthetic wastewater for thermophilic hydrogen production.

**Author Contributions:** Conceptualisation, N.S.J., S.S.M., and N.F.D.R.; methodology, N.F.D.R. and S.S.M.; validation, N.F.D.R.; investigation, N.F.D.R.; resources, N.S.J.; writing—original draft preparation, N.F.D.R.; visualization, S.F.I.; software, S.I.S.; supervision, N.S.J., N.A., and M.F.I.; project administration, N.S.J. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the Ministry of Education Malaysia (MOE) under the Fundamental Research Grant Scheme (FRGS) through the vot code of 5540208 (Ref: FRGS/1/201/STG05/UPM/02/28) towards the success of this study.

**Acknowledgments:** These authors would like to express their gratitude to all the laboratory staff at the Department of Chemical and Environmental Engineering, Universiti Putra Malaysia, for assisting this research.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


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