polymers
Article
Biosynthesis of Poly-ß-Hydroxybutyrate (PHB) from Different
Bacterial Strains Grown on Alternative Cheap Carbon Sources
Sherif M. El-Kadi 1 , Mohssen Elbagory 2,3 , Hassan A. H. EL-Zawawy 4 , Hossam F. A. EL-Shaer 5 ,
Adel A. Shoukry 4 , Sahar El-Nahrawy 3 , Alaa El-Dein Omara 3, * and Dina Fathi Ismail Ali 6
1
2
3
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6
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Citation: El-Kadi, S.M.; Elbagory, M.;
EL-Zawawy, H.A.H.; EL-Shaer, H.F.A.;
Shoukry, A.A.; El-Nahrawy, S.; Omara,
A.E.-D.; Ali, D.F.I. Biosynthesis of
Poly-ß-Hydroxybutyrate (PHB) from
Different Bacterial Strains Grown on
Alternative Cheap Carbon Sources.
Polymers 2021, 13, 3801. https://
doi.org/10.3390/polym13213801
Academic Editor: Edina Rusen
Received: 4 October 2021
Accepted: 29 October 2021
Published: 3 November 2021
Publisher’s Note: MDPI stays neutral
with regard to jurisdictional claims in
Agricultural Microbiology Department, Faculty of Agriculture, Damietta University, Damietta 22052, Egypt;
sherifelkadi@du.edu.eg
Department of Biology, Faculty of Science and Arts, King Khalid University,
Mohail Assir 61321, Saudi Arabia; mhmohammad@kku.edu.sa
Department of Microbiology, Soils, Water and Environment Research Institute, Agricultural Research Center,
Giza 12112, Egypt; sahar.elnahrawy@yahoo.com
Botany Department (Microbiology), Faculty of Agriculture, Al-Azhar University, Cairo 11651, Egypt;
zawawy.hassan@gmail.com (H.A.H.E.-Z.); alshokry17@azhar.edu.eg (A.A.S.)
Botany Department (Genetics), Faculty of Agriculture, Al-Azhar University, Cairo 11651, Egypt;
hosamelshaer805@azhar.edu.eg
Agricultural Microbiology Department, Faculty of Agriculture, Mansoura University, Mansoura 35516, Egypt;
dfali@mans.edu.eg
Correspondence: alaa.ahmed@arc.sci.eg or alaa.omara@yahoo.com
Abstract: Thirty bacterial isolates were tested on three different media for Poly-ß-hydroxybutyrate
(PHB) production. The best bacterial isolates for producing PHB were screened and identified based
on molecular biology; then, using three different alternative carbon sources (dried whey, sugar
beet molasses and date molasses), physical properties were evaluated by Infrared (IR) spectrometry
and Gas chromatography–mass spectrometry (GC-MS/MS) analysis. Our results showed that
the best isolates identified based on molecular biology were Bacillus paramycoides MCCC 1A04098,
Azotobacter salinestris NBRC 102611 and Brevundimonas naejangsanensis BIO-TAS2-2. The addition
of sugar beet molasses to the medium of A. salinestris increased the cell dry weight (CDW), PHB
concentration, PHB% and conversion coefficient (4.97 g/L, 1.56 g/L, 31.38% and 23.92%, respectively).
The correlation coefficient values between PHB g/L and CDW g/L varied between very strong and
moderate positive correlation. IR of the produced PHB from B. paramycoides and A. salinestris showed
similar bands which confirmed the presence of PHB; however, B. naejangsanensis showed weak bands,
indicating lower PHB concentration. The chemical composition obtained showed that the GC-MS
of the PHB extracted represents 2, 4-ditert-butylphenol for B. paramycoides and isopropyl ester of 2butenoic acid for both of A. salinestris and Brevundimonas naejangsanensis. Therefore, PHB produced by
microorganisms can be considered a biodegradable polyester, and represents a promising technique
for the development of eco-friendly and fully biodegradable plastics.
Keywords: agro-industrial waste; GC-MS/MS spectrometry; IR spectrometry; PHB
published maps and institutional affiliations.
1. Introduction
Copyright: © 2021 by the authors.
Licensee MDPI, Basel, Switzerland.
This article is an open access article
distributed under the terms and
conditions of the Creative Commons
Attribution (CC BY) license (https://
creativecommons.org/licenses/by/
4.0/).
Since petroleum-based plastics have become a major cause of environmental pollution,
degradable plastics have attracted a great deal of attention worldwide for their unique
properties, such as thermoplasticity. However, production of these vital plastics is limited
due to their high cost of production. Hence, studies looking for bacterial strains with better
yields and improving their culture conditions are of the greatest importance in order to
reduce the costs of PHB production [1]. The slow growth of the PHB market since the 1980s
is due to the high cost of production for the amount of PHB produced. Currently, only
about 1% of plastic materials on the market are biodegradable, and with the increase in
environmental pollution, there is a need to increase the production of PHB [2]. PHB and
Polymers 2021, 13, 3801. https://doi.org/10.3390/polym13213801
https://www.mdpi.com/journal/polymers
Polymers 2021, 13, 3801
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different biodegradable polyesters are promising tools for the development of eco-friendly
and fully biodegradable plastics. PHBs are a class of biopolymers produced by different
microbial species. Unlike plastics derived from petrochemicals, PHB is biodegradable
and biocompatible in nature. PHBs have advanced applications in the medical sector
(surgical pins and sutures, wound dressings, bone replacements), the packaging industry
(packaging films, bags, cosmetics containers, shampoo bottles), and agriculture (plastic
mulch, as a carrier for long-term doses of herbicides, fungicides and insecticides) [3–6].
PHB is produced using various raw materials such as agricultural and food industry
wastes, which contributes to its economic feasibility. Research is currently concerned with
reducing the cost of production [7]. The use of food industrial byproducts in the production
of PHB is considered one of the growing areas of modern biotechnology; these products
can be used as a nutrient-rich medium at no additional cost. Whey and molasses are very
important byproducts that are rich in nutrients and which bacteria can convert into PHB [8].
The use of molasses as a potential low-cost substrate for PHB biopolymer production has
been studied by [9]. The use of molasses, which is a cheap substrate available in Egypt,
may help reduce the cost of producing such biopolyesters [3]. Beet molasses has been
shown to be an excellent starting material for PHB production by Azotobacter vinelandii. The
substrate cost for PHB production from beet molasses in fed-batch culture was one-third of
that using glucose [10]. The results obtained by [11] indicate that by using Bacillus subtilis
RS1, agro-industry wastes such as sugarcane molasses can be used as an inexpensive
substrate for PHB production. Date molasses and date seed oil have also been used as
the sole carbon source for PHB biosynthesis [12]. Processes are currently underway to
improve PHB production via genetic modification of some strains in order to develop
desirable and suitable properties for some diverse applications [7]. Pavan [13] aimed
to conduct an economic assessment of PHB production process using molasses. The
alternative sources showed equally low production costs (USD 4.28 per kg) and better
economic indicators among the evaluated scenarios. In recent years, using renewable
raw materials for PHB production as degradable bioplastics has been a good method of
environment consideration. Low-cost carbon substrates have the potential to reduce PHB
production cost and increase sustainability [14]. According to [11], the global PHB market
is expected to reach USD 93.5 million by 2021. In the Middle East and Arab countries, Date
palm (Pheonix dactylifera L.) is an important commercial crop. Egypt is one of the largest
date producers among Arab and world countries. Egyptian date production in 2020 was
1.9 million tons, which represented 21.2% of world production. Date soft types are highly
perishable and easily attacked by microbial enzymes; thus, date and date molasses are
considered good materials for fermentation processes [15–17]. The main reason for the low
prevalence of bioplastic products is the high cost of production.
Therefore, this study aims to produce PHB using efficient PHB-producing bacteria on
different available alternative carbon sources (whey, sugar beet molasses and date molasses)
and to study their properties by infrared (IR) spectroscopy and gas chromatography GCMS/MS mass spectrometry analysis.
2. Materials and Methods
2.1. Bacterial Isolation
Four different soil samples were obtained from Damietta and Dakahlia Governorates
(New Damietta city, Kafer El-Batekh, Kafer Saad, and Kalabshou), Egypt, were used for
bacterial isolation. From these soil samples, 30 bacterial isolates were randomly isolated on
three different selective media (10 for each medium): medium (A) [18], medium (B) [19] and
medium (C) [20], for PHB production. These isolates were identified in the Agricultural
Microbiology Department, Faculty of Agriculture, Damietta University, Damietta, Egypt.
Morphological and physiological characteristics of bacterial isolates such as shape, Gram
stain, spore stain, capsule stain and motility were microscopically studied, as were indole
and catalase tests, starch hydrolysis and casein hydrolysis [21–23].
Polymers 2021, 13, 3801
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Isolates obtained from each of the three previously-mentioned media (10 randomly
picked from each medium) were maintained on slants of Nutrient agar medium [24],
Modified Ashby’s medium [25], and Nutrient-rich medium [26], incubated at 5 ◦ C, and
sub-cultured monthly.
2.2. Efficacy of PHB-Producing Isolates
2.2.1. Cultivation Media
A cultivation process was implemented so that isolates from each medium could be
cultivated on the same isolation media for PHB production. Carbon and nitrogen sources
for each medium were sterilized separately and were aseptically added before inoculation
and after cooling of each medium.
For medium (A) [18], 20 g/L glucose (carbon source) and 2.0 g/L ammonium acetate
(nitrogen source) were added to the sterilized medium. The pH was adjusted to 7. For to
medium (B), 10 g/L fructose was added as a carbon source [19] and 1.2 g/L ammonium
acetate (nitrogen source) was added to the sterilized medium; the pH was adjusted to
7.2. Finally, 30 g/L sucrose was used as a carbon source for medium (C) [20], and 1 g/L
ammonium sulfate was added to the sterilized medium. The pH was adjusted to 6.8.
2.2.2. Cultivation System
Batch cultivation on conical flasks was used in this experiment. Twenty ml of each
medium was prepared from maintenance cultures to be used as an inoculum. Incubation
was performed in an incubator shaker at 200 rpm at 30 ◦ C for 48 h. The inoculum was
aseptically transferred to 50 mL of the same medium and incubated under the same
conditions for the same period mentioned above [27]. All experiments were conducted in
three replications.
2.2.3. Biomass Determination
A 50 mL sample of each culture was centrifuged at 5000× g for 10 min. Cell pellets
were washed twice with sterile distilled water. The precipitated biomass was dried at
100 ◦ C for 24 h or until a constant weight was reached. This value was used for determination of cell dry weight (CDW) [28].
2.3. PHB Determination
Ten ml of chloroform was added to the dried cell biomass obtained from the previous
centrifuged cultures and incubated at 70 ◦ C for 10 min, then centrifuged at 5000× g for
10 min. The resulting solution was separated and collected, and the chloroform was
allowed to evaporate in order to obtain PHBs for further assessment. Then, the precipitate
was dried at 100 ◦ C for 24 h or until a constant weight was reached. This value was
used for PHB–free cell dry weight determination [19]. The difference in weight between
the previous two values was considered as PHB production (g/50 mL); this value was
calculated by [29,30], as g/L:
PHB (%) = PHB weight (g/L) × 100/total cell dry weight (g/L)
Conversion coefficient (%) = PHB weight (g/L) × 100/utilized sugar weight (g/L)
2.4. Determination of Utilized Sugars and Nitrogen
Total carbohydrates were determined as glucose [31]. The total sugars remaining
in the culture broth were estimated, and by subtracting this value from the amount of
total sugars in the medium composition at the beginning of the experiment, the consumed
sugars or utilized sugars were calculated. Total nitrogen was measured according to the
Kjeldahl digestion method [32].
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2.5. The Relationship between PHB Production, CDW and Utilized Sugar (g/L)
The correlation coefficient values (r) was calculated according the equation given
in [33]. The correlation between PHB g/L and CDW g/L was calculated where X is the
value of PHB g/L, X is the arithmetical mean of PHB g/L, Y is the value of CDW g/L, and
Y is the arithmetical mean of the CDW g/L values. The value of CDW was replaced with
the value of utilized sugar when the correlation coefficient was calculated between PHB
g/L and utilized sugar.
∑ X−X Y−Y
q
r= q
2
2
∑ X−X
∑ Y−Y
If the value of the correlation coefficient (r) = +1, there is a perfect positive correlation;
if (r) = −1, a perfect negative correlation; if r = 0, there is no correlation. If the value of the
correlation coefficient is between r = 0.0–0.2, 0.2–0.4, 0.4–0.6, 0.6–0.8 or 0.8–1.0, this means a
very weak correlation, weak correlation, moderate correlation, strong correlation or very
strong correlation, respectively [33].
2.6. Identification of Bacterial Isolates Based on Molecular Biology
The highest PHB-producing isolates were inoculated in suitable media and incubated
overnight at 37 ◦ C. The resulting bacterial suspension was pelleted at 10,000 rpm for
5 min and genomic DNA was extracted using a Pure Link genomic DNA mini kit. PCR
amplification of 16s rDNA was performed using the isolated DNA. The PCR conditions
were set as follows: initial denaturation at 95 ◦ C for 5 min, followed by 25 cycles of
denaturation at 95 ◦ C for 40 s, annealing at 55 ◦ C for 2 min and primer extension at 72 ◦ C
for 1 min, ending with the final elongation step at 72 ◦ C for 7 min. PCR products were
gel purified and sent for sequencing with 16S rRNA primer. The obtained sequences were
trimmed to obtain a sequence with ladder 1–1.5 Kb DNA. Then, the sequences were BLAST
search analyzed on the (www.ncbi.nlm.nih.gov, accessed on 1 September 2021) to identify
the isolate [34].
2.7. PHB Production Using Cheap Alternative Carbon Sources
Three different alternative carbon sources were used; dried whey and date molasses
were purchased from Damietta City market, Damietta City, Egypt, and sugar beet molasses
was purchased from the Sugar Beet Factory Kalabsho, Belkas, Dakahlia Governorate, Egypt.
Molasses was pretreated according to the method in [35]. Total sugars and total nitrogen of
all sources were determined. The values of carbon and nitrogen were considered during
preparation of the cultivation media which already contain these sources.
2.8. The Physical Properties of PHB Polymer
IR was used to confirm the formation of PHB using a MATTISON 5000 FTIR spectrometer at Mansoura University, Faculty of Pharmacy, Chemistry Department, Spectral
Analysis Unit. A 50 mm mortar and pestle were used. Hydraulic presses were performed
using a PIKE die press. A SPECAC press instrument was used for preparation of the KBr
sample discs of investigated drugs, pressing under a 7 mm die at a pressure of 2 tons
for 1 min. A SHIMADZU CORPORATION balance BL-220H (Kyoto, Japan) was utilized
throughout the work. IR spectra of the PHB polymer were determined at 2 cm−1 spectral
resolution by using a MATTSON 5000 FTIR spectrometer. An excitation wavelength at
3900 nm was provided and the laser power at the sample position was typically 500 nm.
Spectra were obtained with a spectral resolution of 4 cm−1 , and 1024 [36].
For GC-MS/MS analysis, the chemical composition was obtained after extraction
and derivatized using BSTFA reagent. The derivatization product MTBSTFA-HB was
analyzed on a Finnigan GCQ (Finnigan MAT, Austin, TX, USA), which is an ion trap MS
with external ionization. A 30 m fused silica capillary column DB-5MS, I.D. 0.25 mm,
film thickness 0.25 mm (J&W Scientific, Folsom, CA, USA) was used for separation. The
samples were injected in splitless mode. The injection temperature was 250 ◦ C, temperature
Polymers 2021, 13, 3801
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of ion source 200 ◦ C and transfer line 275 ◦ C. Helium velocity was 40 cm s−1 . The oven
temperature, initially at 60 ◦ C, was gradually increased (20 ◦ C min−1 ) to 250 ◦ C and held at
this temperature for 15.5 min. The MS/MS detection consisted of a sequence of three steps:
(i) isolation of the “parent” ion of the detected compounds (immediate rejection of other
ions except the “parent” ion); (ii) fragmentation of the “parent” ion; and (iii) analysis of the
fragmentation products. These steps were realized using GCQ MS/MS software, version
2.0. The MS/MS measurements were performed at a collision energy of 0.8 V for parent
ions of m/z 275 (for the analyte) and m/z 331 (for the internal standard). The product
spectra in the mass range of m/z 100–331 were scanned. The retention time of the analyte
was 8.34 ± 0.04 min and of the internal standard 10.2 ± 0.04 min [37].
2.9. Statistical Analysis
Data obtained throughout this study were analyzed by computer-assisted one-way
ANOVA, using the software package stat graphics version 5.0 (costat). Least significance
differences (LSDs) were calculated at level of significance p < 0.05 [38].
3. Results
3.1. Morphological and Physiological Characteristics of the Bacterial Isolates
Thirty bacterial isolates (10 isolates for each medium) were randomly isolated on three
different selective media, medium (A), medium (B), and medium (C). Some of morphological and physiological characteristics of bacterial isolates were obtained. In addition,
10 bacterial isolates which were isolated on medium (A) were bacilli, Gram positive, spore
positive, capsule positive and negative, and motile, and the indole test, catalase test, starch
hydrolysis and casein hydrolysis were positive (Table S1, Supplementary Materials). From
these characteristics, all isolates seemed to be Bacillus spp. [21]. Finally, 10 bacterial isolates
which were isolated on medium (B) were diplococcic, Gram negative, non-spore, capsule
positive, and motile and non-motile, and the indole test, catalase test, starch hydrolysis
and casein hydrolysis were positive (Table S2, Supplementary Materials). From these
characteristics, all isolates seemed to be Azotobacter spp. [21].
Bacterial isolates which were isolated on medium (C) were bacilli, cocci and short rods.
Gram stain, spore stain, and capsule were positive and negative. Motile and none motile,
indole test, catalase test, starch hydrolysis and casein hydrolysis were positive (Table S3,
Supplementary Materials).
3.2. Screening the Best PHB-Producing Bacterial Isolates
The results presented in Table 1 show that all tested isolates cultivated on medium
(A) produced PHB. The CDW varied from 0.98 to 2.99 g/L. PHB production ranged from
0.10 to 0.62 g/L. PHB% ranged between 6.84 and 27.55%. Sugar utilization ranged from
14.32 g/L to 18.51 g/L. The values of the conversion coefficient ranged between 0.62%
and 3.90%. Significant differences were observed between bacterial isolates in most cases.
Generally, the best isolate which yielded the highest PHB production with reasonable sugar
utilization was No. A6. Therefore, this isolate was chosen and used in the subsequent
experiments. Isolates No. A1, A2 and A6 also produced more than 20% PHB.
In addition, all tested isolates grown on medium (B) also produced PHB (Table 1). The
highest values of cell dry weight, PHB production, PHB%, sugar utilization and conversion
coefficient were 4.00 g/L, 0.93 g/L, 28.51%, 20.93 g/L, and 6.17%, respectively. However,
the lowest values of the above parameters were 2.56 g/L, 0.73 g/L, 19.50%, 15.07 g/L,
and 2.86%, respectively. It was observed that there were significant differences between
the bacterial isolates; thus, the best isolate with the highest value of PHB (0.93 g/L) was
No. P3. This isolate was therefore chosen for identification based on molecular biology
and used in the experiments on alternative carbon sources. PHB was produced from all
isolates grown on medium (C) (Table 1). The CDW values varied from 0.79 to 3.1 g/L.
PHB production ranged from 0.08 to 0.73 g/L, and PHB% ranged between 6.04 and 23.67%.
Sugar utilization ranged from 6.09 g/L to 8.17. The values of the conversion coefficient
Polymers 2021, 13, 3801
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ranged between 1.00% and 8.93%. Generally, the best isolate which yielded the highest
PHB production (0.73 g/L) with reasonable sugar utilization was No. W8. It was noted
that there were significant differences between the bacterial isolates. Therefore, this isolate
was chosen and used in the subsequent experiments.
Table 1. Screening of different bacterial isolates to produce PHB on different media.
Isolate
Number
C.D.W (g/L)
PHB (g/L)
PHB (%)
Utilized
Sugar (g/L)
Conversion
Coefficient (%)
Medium A
A1
A2
A3
A4
A5
A6
A7
A8
A9
A10
2.99 a
2.52 b
1.27 f
1.69 d
1.17 fg
2.25 c
0.98 h
1.46 e
1.03 gh
1.58 de
0.61 b
0.51 b
0.11 de
0.15 de
0.13 de
0.62 a
0.17 d
0.10 e
0.12 de
0.29 c
20.40 c
20.23 b
8.66 h
8.87 h
11.11 g
27.55 a
17.34 e
6.84 i
11.65 f
18.35 d
17.03 c
15.36 d
16.00 d
14.32 e
18.51 a
15.88 d
16.85 c
15.94 d
17.77 b
15.80 d
3.58 b
3.32 b
0.68 e
1.04 d
0.70 e
3.90 a
1.00 d
0.62 e
0.67 e
1.83 b
LSD 0.05
0.145
0.042
0.404
0.486
0.140
Medium B
P1
P2
P3
P4
P5
P6
P7
P8
P9
P10
3.61 b
3.93 a
3.47 bc
3.30 c
4.00 a
3.62 b
2.78 d
2.56 e
3.88 a
3.40 bc
0.85 b
0.83 b
0.93 a
0.83 b
0.78 bc
0.78 bc
0.74 c
0.73 c
0.82 b
0.84 b
23.54 c
21.11 d
26.80 ab
25.14 bc
19.50 d
21.54 d
26.62 ab
28.51 a
21.12 d
24.75 bc
15.34 e
19.58 c
15.07 e
15.19 e
15.44 e
17.32 d
19.17 c
20.93 b
19.40 c
29.33 a
5.54 b
4.23 de
6.17 a
5.46 b
5.05 c
4.50 d
3.86 e
3.48 f
4.22 de
2.86 g
LSD 0.05
0.165
0.053
1.571
0.455
0.365
d
ef
Medium C
W1
W2
W3
W4
W5
W6
W7
W8
W9
W10
1.37
2.04 b
1.85 c
1.15 e
1.36 d
1.98 b
1.82 c
3.10 a
0.79 f
1.33 d
0.10
0.28 b
0.17 d
0.17 d
0.21 c
0.16 d
0.13 de
0.73 a
0.11 ef
0.08 f
6.6 d
13.72 b
9.37 c
15.40 b
14.73 b
8.25 cd
7.68 cd
23.67 a
14.79 b
6.04 d
7.94 a
6.82 e
7.71 b
7.39 c
7.10 d
6.09 f
7.65 b
8.17 a
7.44 c
7.96 a
1.25 ef
4.10 b
2.20 d
2.30 d
2.95 c
2.62 cd
1.69 e
8.93 a
1.47 ef
1.00 f
LSD 0.05
0.121
0.032
1.748
0.203
0.433
C.D.W: cell dry weight; PHB: Poly-ß-hydroxybutyrate; A1–10: symbol for unknown bacterial isolate grown on
medium (A) [15]; P1–10: symbol for unknown bacterial isolate grown on medium (B) [16]; W1–10: symbol for
unknown bacterial isolate grown on medium (C) [17]. Means followed by different letters indicate significant
differences between treatments according to Duncan’s test (p < 0.05).
3.3. Identification of the Best Bacterial Isolates Based on Molecular Method
Isolates No. A6, P3 and W8 (which were the best isolates for PHB production in the
previous experiments) were identified according to the Polymerase Chain Reaction (PCR)
method by Sigma Company (Cairo, Egypt) [34]. Isolate No. A6 was identified by the
Polymers 2021, 13, 3801
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16S ribosomal RNA gene and the partial sequence Query1 GAGCTTGCTCTTATGAAGTTAGCGGCGGACGGGTGAGTAACACGTGGGTAACCTGCCCAT 60.
The sequences of this isolate were accessed through a database (www.ncbi.nlm.nih.
gov, accessed on 1 September 2021) using the accession number, and it was found to belong
to the Bacillus paramycoides strain MCCC 1A04098 (Figure 1A). Isolate No. P3 was identified
using the same method, by the partial sequence Query 1 AATACCCTGCAGTCTTGACGTTACCGGCAGAATAAGCACCGGCTAACTTCGTGCCAGCAG 60. It was identified as the
Azotobacter salinestris strain NBRC 102611 (Figure 1B). Isolate No. W8 was also identified
using the same method, by the partial sequence Query 1 GAATAACTCAGGGAAACTTGTGCTAATACCGAATGTGCCCTTTGGGGGAAAGATTTATCG 60. It was identified as
Brevundimonas naejangsanensis strain BIO-TAS2-2 (Figure 1C).
3.4. PHB Production from Cheap Alternative Carbon Sources
The values of carbon and nitrogen were considered during preparation of the cultivation media, which already contained these sources. In dried whey, sugar beet molasses
and date molasses, the carbon contents were 70%, 51%, and 37%, respectively, and the
nitrogen contents were 15%, 13%, and 14.1%, respectively. Table 2 shows the effect on PHB
production from B. paramycoides strain MCCC 1A04098 of replacing glucose in medium
(A) with the three cheap alternative carbon sources (dried whey, treated sugar beet molasses, and treated date molasses). The addition of treated sugar beet molasses was the
most economically suitable, as it increased PHB production (0.98 g/L) and the conversion
coefficient (6.5%). Furthermore, the utilized sugar (15.11 g/L) was moderate and slightly
lower than the control value (15.88 g/L). On the other hand, dried whey yielded the lowest
PHB production (0.51 g/L), even lower than the control. The values of treated date molasses showed the same pattern as treated sugar beet molasses, except for sugar utilization
which was the highest among all carbon sources (16.40 g/L). It was observed that there
were significant differences between bacterial strains. In the present study, the maximum
production of PHB and CDW by B. paramycoides was 0.98 g/L and 2.84 g/L from sugar
beet molasses, respectively.
A
Figure 1. Cont.
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B
C
Figure 1. Genotype tree of Bacillus paramycoides MCCC 1A04098 (A), Azotobacter salinestris NBRC
102611 (B) and Brevundimonas naejangsanensis BIO-TAS2-2 (C).
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Table 2. Effect of cheap alternatives carbon sources on PHB production by different strains of Bacillus paramycoides,
Azotobacter salinestris and Brevundimonas naejangsanensis.
Alternatives Carbon
Sources
C.D.W (g/L)
PHB (g/L)
PHB (%)
Utilized Sugar (g/L)
Conversion
Coefficient (%)
Bacillus paramycoidesstrain MCCC 1A04098
Dried whey
sugar beet molasses
Date molasses
(Control, glucose)
1.56
2.84 a
2.67 b
(2.25 c )
0.51 d
0.98 a
0.88 b
(0.62 c )
33.81 a
34.50 a
32.86 a
(27.55 b )
13.74 b
15.11 ab
16.40 a
(15.88 a )
3.71 c
6.50 a
5.36 b
(3.90 c )
LSD 0.05
0.123
0.079
4.499
1.391
0.804
d
Azotobacter salinestris strain NBRC 102611
Dried whey
Sugar beet molasses
Date molasses
(Control, fructose)
2.47 d
4.97 a
4.61 b
(3.10 c )
0.66 d
1.56 a
1.36 b
(0.73 c )
26.51 bc
31.39 a
29.45 ab
(23.67 c )
5.15 d
6.52 c
6.83 b
(8.17 a )
12.81 c
23.92 a
19.91 b
(8.93 d )
LSD 0.05
0.227
0.047
2.951
0.220
0.984
Brevundimonas naejangsanensis strain BIO-TAS2-2
d
Dried whey
Sugar beet molasses
Date molasses
(Control, sucrose)
1.63
3.57 a
3.19 c
(3.47 b )
0.64 d
1.36 b
1.50 a
(0.93 c )
39.27 b
40.36 b
47.14 a
(26.80 c )
11.59 d
16.71 a
16.03 b
(15.07 c )
5.51 b
9.73 a
9.35 a
(6.17 b )
LSD 0.05
0.161
0.069
3.537
0.284
0.657
Means followed by different letters indicate significant differences between treatments according to Duncan’s test (p < 0.05).
Fructose was replaced with alternative carbon sources in medium (B) for PHB production using the A. salinestris strain NBRC 102611. Table 2 showed that the use of treated beet
molasses yielded the highest PHB production (1.56 g/L) and highest conversion coefficient
(23.92%), with moderate sugar utilization (6.52 g/L) in comparison with the control values.
Dried whey yielded the lowest PHB production (0.66 g/L). It was observed that there were
significant differences between the bacterial isolates.
Sucrose was replaced with alternative carbon sources in medium (C) for PHB production from the Brevundimonas naejangsanensis strain BIO-TAS2-2. Table 2 showed that the
use of treated date molasses yielded the highest PHB production (1.5 g/L) and the highest
conversion coefficient (9.35%), with moderate sugar utilization (16.03 g/L) in comparison
with the control values. Dried whey yielded the lowest PHB production (0.64 g/L). It was
observed that there were significant differences between the bacterial isolates.
3.5. The Correlation Coefficient Value between PHB Production, CDW and Utilized Sugar (g/L) of
All Tested Isolates and Strains
The correlation coefficient was calculated according the equation in [33]. Statistical
analysis indicated there was a very strong positive correlation (r = 0.987) between PHB
g/L and CDW g/L in the A. salinestris strain and a strong positive correlation (r = 0.457) in
the isolates grown on medium (B). A very weak negative correlation (r = −0.161) between
PHB g/L and utilized sugar g/L in the isolates grown on medium (B) was observed, with
the highest value between them being r = 0.897 (a very strong positive correlation) in the
case of B. naejangsanensis (Table 3).
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Table 3. The correlation analysis between PHB, dry cell weight and utilized sugar.
Strains or Isolates
Obtained From
Correlation Coefficient Analysis (r Value)
Correlation between PHB
g/L and CDW (g/L)
Correlation between PHB
g/L and Utilized Sugar (g/L)
0.901
0.958
0.457
0.987
0.840
−0.012
0.527
−0.161
0.051
0.239
0.724
0.897
Medium (A)
Bacillus paramycoides
Medium (B)
Azotobacter salinestris
Medium (C)
Brevundimonas
naejangsanensis
Medium (A) [15]; Medium (B) [16]; Medium (C) [17].
3.6. The Physical Properties of PHB Polymer
3.6.1. Fourier Transform Infrared Spectroscopy (FTIR)
Poly ß-hydroxybutyrate (PHB) Polymer extracted from B. paramycoides was characterized using IR spectra in the range of 600–4000 cm−1 , as displayed in Figure 2a. The IR
spectra of PHB produced show obvious peaks at 1262 and 1725 cm−1 , corresponding to
specific rotations around carbon atoms specific to –CH and C=O stretching of the ester
group present in the molecular chain [36]. Strong peaks at 1034 and 1097 indicate the
presence of C-O stretching. Additionally, the absorption bands at 2926 and 2963 cm−1
are due to C–H stretching vibrations of methyl and methylene groups, which confirms
PHB formation. PHB extracted from A. salinestris, as shown in Figure 2b, is similar to that
extracted from B. paramycoides. Furthermore, the IR spectra of the extract from A. salinestris
display similar bands at 1032, 1098 1262, 1725, 2925 and 2963 cm−1 which confirm the
presence of PHB, as shown in Figure 2b. On the other hand, the IR spectra of PHB extracted
from Brevundimonas naejangsanensis illustrate weak bands at 1264 and 1748 cm−1 , indicating
lower PHB concentration in comparison with the other obtained extracts, as shown in
Figure 2c. In addition, all peaks at 3400 to 3600 cm−1 correspond to specific amino groups.
3.6.2. Gas Chromatography–Mass Spectrometry (GC-MS) Analysis
Chemical composition was obtained after extraction and derivatized using BSTFA
reagent. Figure 3a shows a peak at retention time 15.91 min, which represents Polyhydroxybutyrate (PHB) from B. paramycoides. The molecular ion peak in the mass spectrum at m/z
104 agrees well with the molecular formula for PHB. In addition, Figure 3b shows a peak
at 20.08 min, which represents 2, 4-ditert-butylphenol. The area under the peak denotes
the PHB content, which was determined to be 4.85%.
Figure 4a shows the GC-MS chromatogram of the PHB extracted from A. salinestris;
the peak at retention time 16.07 min represents the isopropyl ester of 2-butenoic acid
(Figure 4b), confirming the present of Polyhydroxybutyrate (PHB). The area under the
peak denotes the PHB content, which was determined to be 10.73% (Figure 4b). Figure 5a
shows the GC-MS chromatogram of the PHB extracted from Brevundimonas naejangsanensis,
showing a peak at retention time 16.07 min which represents the isopropyl ester of 2butenoic acid (Figure 5b), confirming the presence of Polyhydroxybutyrate (PHB). The area
under the peak denoting the PHB content was determined to be 0.85%, with very low mass
spectrum matching.
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Figure 2. Cont.
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Figure 2. IR spectrum of PHB extracted from B. paramycoides (a), A. salinestris (b), and Brevundimonas naejangsanensis (c).
(a)
Figure 3. Cont.
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(b)
Figure 3. (a) Total GC-MS chromatogram of PHB extracted from Bacillus paramycoides. (b): The mass spectrum of isopropyl
ester of 2-butenoic acid of PHB extracted from Bacillus paramycoides.
(a)
(b)
Figure 4. (a): Total GC-MS chromatogram of PHB extracted from Azotobacter salinestris. (b): The mass spectrum of isopropyl
ester of 2-butenoic acid of PHB extracted from Azotobacter salinestris.
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(a)
(b)
Figure 5. (a): Total GC-MS chromatogram of PHB extracted from Brevundimonas naejangsanensis. (b): The mass spectrum of
isopropyl ester of 2-butenoic acid of PHB extracted from Brevundimonas naejangsanensis.
4. Discussion
4.1. Screening the Best PHB-Producing Bacterial Isolates
Significant differences were observed between the bacterial isolates in most cases.
Generally, the best isolate which yielded the highest PHB production with reasonable sugar
utilization was No. A6 (Table 1). Isolates No. A1, A2 and A5 produced more than 20%
PHB. These results were similar to the results obtained by [39], who isolated three bacteria
from the roots of beans (Vicia faba) grown in Qalyubia Governorate, Egypt to produce PHB
using the flask culture technique. These isolates collected more than 20% of PHB, and were
identified as Pseudomonas fluorescens S48, B. megaterium 7A and B. megaterium UBF19. Four
isolates (A1, A2, A6 and A10) were higher than those found by [18], who reported that the
highest value of PHB in B. megaterium was 0.27 g/L. Furthermore, all of these isolates were
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higher than [40], who produced PHB by batch fermentation of B. subtilis from glucose and
reached only 0.077 g/L.
In addition, all tested isolates grown on medium (B) produced PHB (Table 1). It
was observed that there were significant differences between bacterial isolates. Thus,
the best isolate with the highest value of PHB (0.93 g/L) was No. P3. Multiple previous studies [19,41] produced PHB from A. chroococcum in amounts ranging from 2.0 to
3.0 g/L, higher than in the current study; however, [10] produced only about 1.0 g/L from
A. vinelandii, lower than this study. In [42], the production of PHB was studied using the
shaking culture method with A. chroococcum resulting in a PHB% of 46.80%. On the other
hand, PHB was produced from all isolates grown on medium (C) in the present study
(Table 1).
4.2. Identification of the Best Bacterial Isolates Based on Molecular Method
Isolates No. A6, P3 and W8 were identified according to the Polymerase Chain
Reaction (PCR) method. The sequence of the A6 isolate was accessed using the accession
number and it was found to belong to the Bacillus paramycoides strain MCCC 1A04098
(Figure 1A). Although several reports have produced PHB from Bacillus sp., such as
B. amyloliquefaciens, B. aryabhattai, B. brevis, B. cereus, B. circulans, B. coagulans, B. firmus,
B. laterosporus, B. licheniformis, B. macerans, B. megaterium, B. sphaericus, B. subtilis, and
B. thuringiensis [1,14,43], and a few researchers [44–48] have used B. mycoides for producing
PHB, to the best of our knowledge only this study has produced PHB from B. paramycoides.
In addition, the isolate No. P3 was identified as Azotobacter salinestris strain NBRC
102611 (Figure 1B). Three species of the genus Azotobacter sp. have commonly appeared
in the literature on the production of PHB, including A. chroococcum [19,28,41,42,49],
A. vinelandii [50–53], and A. beijerinckii [54–56]; however, only one used A. salinestris for
PHB production [57] before this study. Isolate No. W8 was identified as the Brevundimonas
naejangsanensis strain BIO-TAS2-2 (Figure 1C). Only one study has used Brevundimonas
naejangsanensis for PHB production [58] prior to the current study. This study is also the
first to report the newly isolated bacterial strain designated as Bacillus paramycoides strain
MCCC 1A04098 as potential source for PHB production.
4.3. PHB Production from Cheap Alternative Carbon Sources
Table 2 shows the effect of replacing glucose in medium (A) with three cheap alternative carbon sources, namely dried whey, treated sugar beet molasses, and treated date
molasses, on PHB production from B. paramycoides strain MCCC 1A04098. Generally, in order to decrease the cost of PHB and increase the cell dry weight, PHB concentration, PHB%,
and conversion coefficient, the addition of treated sugar beet molasses in the medium of
B. paramycoides is recommended.
Bacillus sp. is capable of producing hydrolytic enzymes, which can degrade agroindustrial and other wastes that can be utilized as carbon sources for PHB production [43].
These results were similar to [46], where the highest PHB yield was observed in B. mycoides
DFC1 (1.28 g/L) using wheat starch, and lower than [47], where a maximum dry cell weight
of 4.35 g/L was obtained. In addition, these results were higher than those obtained in [59],
where the maximum PHB% obtained by B. subtilis was 13.02% ± 1.67%. As for the attempts
that have been made to reduce the cost of production, there have been many attempts,
such as [1], who reported that PHB production from B. aryabhattai showed good polymer
accumulation in basal medium with glycerol after 48 h of incubation and accumulated
1.79 g/L. Hence, glycerol may be a better option as it is cheaper than glucose and can be
obtained as a byproduct from industries such as biodiesel production. In the same trend,
the current study was better than [3], who studied the effect of different carbon sources on
PHB production by B. megaterium. The highest production of PHB was observed with cane
molasses as the sole carbon source (40.8, mg/L). The same results were obtained by [60],
who reported that sugar beet molasses could be a suitable substrate for the production
of PHB with B. megaterium. Cell dry weight was 16.7 g/L and PHB production was
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0.6 g/L with batch cultivation. Sugarcane molasses was also used as a sole carbon source
by [11,61,62], using B. cereus and B. subtilis to obtain a PHB% of 57.5%, 49.9% and 44.7%,
respectively.
Fructose was replaced with alternative carbon sources in medium (B) for PHB production from A. salinestris strain NBRC 102611 (Table 2). In [52] it was found that there are more
than of 300 bacterial species that can produce PHB, of which only a few have been used
on a commercial scale. Molasses and whey have been reported to be excellent substrates
for PHB production [35,57,63]. Similar results were obtained by [60], who reported that
the use of sugar beet molasses in PHB production led to a two- to three-fold increase in
biomass production compared to the control experiment. In [10] it was reported that the
results obtained with beet molasses were the best compared with other sugars, especially
unrefined sugar; in addition, it was proven that it is not only a source of carbon, but
also contains some other nutrients that may interfere with the synthesis of PHB. Thus, to
decrease the cost of PHB, the addition of sugar beet molasses in the medium of A. salinestris
to increase cell dry weight, PHB concentration, PHB% and conversion coefficient appears
very promising.
Sucrose was replaced with alternative carbon sources in medium (C) for PHB production from Brevundimonas naejangsanensis strain BIO-TAS2-2 (Table 2). Both the current study
and [12] used date molasses as a novel carbon source for B. naejangsanensis and Cupriavidus
necator for PHB production. The current findings were homogeneous with [58], who tested
the ability of B. vesicularis to accumulate PHB and found that with acid-hydrolyzed wood
(sawdust) as the substrate, PHB production was improved. In order to reduce the production cost by 50% and make it as economical as possible, more than 96% of sugars must be
consumed, and the cells must contain more than 90% PHB with a yield not less than 64%
of the DCW.
4.4. The Correlation Coefficient Value between PHB Production, CDW and Utilized Sugar (g/L) of
All Tested Isolates and Strains
Multiple studies [64,65] have found a positive correlation between PHB production
and CDW with r = 0.999, which was similar to our results where r = 0.987 (Table 3). The
available studies on the correlation between PHB production and CDW were insufficient,
with little information available; [64–66] published that there was a positive correlation
between PHB and CDW. On the contrary, Aslim [67], reported there was a low correlation
between CDW and the PHB contents of Lactobacillus cultures. Thus, there is a need for
further study on this point. In this context, [68] reported that the correlation between PHB
synthesis and nitrogen fixation was little discussed in purple non-sulfur bacteria species.
In keeping with this trend, [65,69] studied the correlation between PHB production, pH,
and viscosity, finding that it was decreased or that no correlation could be observed.
However, there are no studies dealing with the correlation between PHB production and
the amount of sugar consumed, which is the most important economic factor affecting
polymer production.
4.5. The Physical Properties of PHB Polymer
4.5.1. Fourier Transform Infrared Spectroscopy (FTIR)
Poly ß-hydroxybutyrate (PHB) Polymer extracted from the studied strains was characterized using IR spectra in the range of 600–4000 cm−1 , as displayed in Figure 2. The
results were obtained by [14], which observed that 1624–1724 cm−1 was associated with the
C=O stretching of the ester carbonyl bond. The bond at 1529 cm−1 was characteristic of the
stretching and deformation vibration of the C-H group, and those at 2960 and 3277 cm−1
were characteristic of the stretching and deformation vibrations of the terminal OH groups.
The PHB polymer functional groups were confirmed as C=O groups by FT-IR spectroscopy.
In addition, analysis of the spectrum of PHB was observed at 3400, 1639, 3018, 2978, 2842,
1216, 1044 and 669–765 δ, respectively, for the (-OH) broad peak, (C=C) double bond, (≡CH) acetylenic bond, (-CH3) methyl group, (-S-H) thiol weak adsorption, (≡C-O-C≡) ether,
Polymers 2021, 13, 3801
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sulfoxide and (F-Cl) haloalkanes [41]. The same results were obtained by [70], who showed
that the bands of PHB between 3300 and 3700 cm−1 were ascribed to O-H groups in the
phenolic and aliphatic structures. The band at 2923 cm−1 was assigned to C-H stretching
in aromatic methoxyl groups, or in methyl and methylene groups of the side chain. The
band at 1645 cm−1 was attributed to unconjugated C=O stretching. The bands centered
at 1508 and 1593 cm−1 were associated with aromatic structural vibrations. The bands
between 1400 and 1450 cm−1 originated from C-H deformation coupled with the vibration
from the aromatic ring. The bands at 1217 and 1128 cm−1 indicated guaiacyl and syringyl
groups, respectively. Moreover, Rathika [11], reported that, the broad transmittance peak
at 3281 cm−1 could be ascribed to the stretching of O–H groups. Peaks observed at 2924
and 2852 cm−1 were associated with the C-H stretching of bonds of methyl (CH3) and
methylene (CH2) groups. The intense absorption peak at 1722 cm−1 was the characteristic
carbonyl (C=O) stretching of ester groups in the extracted PHAs.
The 1H NMR spectrum showed the expected resonances for PHB as demonstrated
by the methine group (-CH-) between 5.22 and 5.28 ppm, a methylene group (-CH2-)
between 2.45 and 2.62 ppm, and the methyl group (-CH3) between 1.26 and 1.28 ppm, as
in standard PHB. Scans of 13C NMR showed peaks at 169.13, 67.62, 40.81 and 19.76 ppm,
which represent the carbonyl carbon (-C-), ester (-O-CH-) group, methylene (-CH2-) and
the methyl (-CH3) groups, as shown in the standard. These results confirm the material as
a homopolymer of 3-hydroxybutyrates, i.e., poly-3-hydroxybutyrate [1].
4.5.2. Gas Chromatography–Mass Spectrometry (GC-MS) Analysis
Our observation in Figures 3–5 were in agreement with [11], who detected PHAs
extracted from B. subtilis RS1 cultured and processed in a medium of sugarcane molasses
using GC–MS and revealed significant peaks at retention times 20.9, 23.1, and 23.8 min,
corresponding to methyl esters of pentadecanoic acid and hexadecanoic acid, respectively.
Furthermore, the results revealed that hexadecanoic acid methyl ester is the predominant
monomer of PHA produced from B. subtilis RS1. In addition, Ramya [14] reported that the
bioplastics produced by B. cereus RBL6 were characterized by GC–MS. B. cereus RBL6 was
reported to produce methyl-3-hydroxy hexadecanoic acid, which belongs to the monomer
chains of the medium-chain-length PHA class.
5. Conclusions
Our study aimed to use cheap and available alternative carbon sources (whey, sugar
beet molasses and date molasses) to produce PHB using Bacillus paramycoides, Azotobacter
salinestris and Brevundimonas naejangsanensis. The results showed that the addition of
sugar beet molasses in the medium of A. salinestris increased the cell dry weight (CDW),
PHB concentration, PHB% and conversion coefficient. In addition, statistical analysis
indicated that the correlation coefficient values between PHB g/L and CDW g/L varied
between r = 0.987 and r = 0.457 (very strong positive correlation and moderate positive
correlation); however, these values were r = −0.161 and r = 0.897 when the correlation
was calculated between PHB g/L and utilized sugar g/L. In addition, the IR of the produced PHB from B. paramycoides and A. salinestris showed similar bands, which confirmed
the presence of PHB; however, B. naejangsanensis showed weak bands. The chemical
composition obtained by GC-MS of the PHB extract represented 2, 4-ditert-butylphenol
for B. paramycoides, and isopropyl ester of 2-butenoic acid for both of A. salinestris and
Brevundimonas naejangsanensis.
Supplementary Materials: The following are available online at https://www.mdpi.com/article/
10.3390/polym13213801/s1: Table S1. Morphological and physiological characteristics of bacterial
isolates on medium (A); Table S2. Morphological and physiological characteristics of bacterial isolates
on medium (B); Table S3. Morphological and physiological characteristics of bacterial isolates on
medium (C).
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Author Contributions: Conceptualization, S.M.E.-K., A.E.-D.O., M.E. and D.F.I.A.; methodology,
S.M.E.-K., A.E.-D.O., S.E.-N., A.A.S., H.F.A.E.-S., H.A.H.E.-Z. and D.F.I.A.; software, S.M.E.-K.,
A.E.-D.O., H.A.H.E.-Z. and D.F.I.A.; validation, S.M.E.-K., A.E.-D.O., M.E. and D.F.I.A.; formal analysis, S.M.E.-K., A.E.-D.O., H.F.A.E.-S., H.A.H.E.-Z. and D.F.I.A.; investigation, S.M.E.-K., A.E.-D.O.,
H.A.H.E.-Z. and D.F.I.A.; resources, S.M.E.-K., A.E.-D.O., H.F.A.E.-S. and H.A.H.E.-Z. and D.F.I.A.;
data curation, S.M.E.-K., A.E.-D.O., S.E.-N., M.E. and D.F.I.A.; writing—original draft preparation,
S.M.E.-K., A.E.-D.O. and D.F.I.A.; writing—review and editing, S.M.E.-K., A.E.-D.O. and D.F.I.A.;
visualization, S.M.E.-K., A.E.-D.O. and D.F.I.A.; supervision, S.M.E.-K., funding acquisition, M.E. All
authors have read and agreed to the published version of the manuscript.
Funding: This research received no external funding.
Institutional Review Board Statement: Not applicable.
Informed Consent Statement: Not applicable.
Data Availability Statement: The data that supports the findings of this study are contained within
the article or supplementary material and available from the corresponding author upon reasonable
request.
Acknowledgments: The authors extend their appreciation to the Deanship of Scientific Research
at King Khalid University for funding this work through the Research Groups Project under grant
number R.G.P. 1/112/42. All the authors are grateful for the support provided by Faculty of
Agriculture, Damietta University, Faculty of Agriculture, Mansoura University and Soils, Water and
Environment Research Institute (SWERI), Agriculture Research Center (ARC), Egypt.
Conflicts of Interest: The authors declare no conflict of interest.
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