growing season (plant cane cycle experiment 2), adapted from [12].

The legume most productive was sunn hemp crotalaria juncea IAC-1 with 10,264 kg ha-1, followed by velvet-bean with 4,391 kg ha-1 and peanuts IAC-Caiapó and IAC-Tatu with 3,177 kg ha-1 and 1,965 kg ha-1, respectively.

There was an increase of Stalk yield of sugarcane in the average of the five cuts, compared to control treatment (Table 16). It can be seen that the effect of planting green manure in the fields of sugarcane promoted reform of benefits in terms of increased productivity of sugarcane, and this is lasting reaching in this case until the fifth cut, and the only treatment that stood out was the rotation of the witness with sunn hemp. Notably, the sunn hemp had higher dry matter production, and this may be a positive influence on growth of sugarcane. After five harvests, sunn hemp crotalaria was the leguminous crop that induced the greatest sugarcane yield, with 30% increase in cane yield and 35% in sugar yield.

C.V.% (plot) = 7.57, CV% (subplots) = 4.20. Means followed by same lower-case letter in rows and capital letters in columns do not differ (Comparisons among means were made according to Tukey test p> 0.05). For statistical analysis the data were transformed into log (x). Adapted from [16].

**Table 16.** Millable stalk yield of sugarcane plants in five consecutive harvests as a function of treatments.

Crop rotation with non-host species of nematodes, when well planned, can be an efficient method for integrated control of nematodes. It is common in some areas, the practice of cultivation of Fabaceae in the period between the destruction of ratoon sugarcane field and planting the new. There are several plants used in systems of crop rotation with sugarcane, the most common are crotalarias, velvet beans, soybeans and peanuts. However, depending on nematode species occurring in the area, some of these cultures may significantly increase the population of these parasites. Thus, the sugarcane crop, can be greatly impaired by increasing the inoculum potential of the nematodes [58, 59].

The peanut IAC-Caiapó and velvet bean were the leguminous crops that resulted in the greater percentage of AM fungus. The lowest population of *Pratylenchus spp*. was found in the treatments with peanut IAC-Tatu and IAC-Caiapó (Figure 3).

Thet peanut IAC-Caiapó showed a minimum of 10 nematodes per 10 g of roots and a maximum of 470, while on the control this variation was from 80 to 2,510, indicating the smaller presence of the nematode in treatments with peanut IAC-Caiapó (Figure 3).

Crop Rotation Biomass and Effects on Sugarcane Yield in Brazil 27

resulted in higher yields of cane in the second cycle. The recovery of 15N- labeled fertilizers by two successive sugarcane crops summed up 19 to 21% of the N applied as sunn hemp and 46 to 49% of the N applied as ammonium sulfate. Very little inorganic N was present in

The sugar content of sugarcane stalks is important because the raw material remuneration takes into account this parameter in Brazil. Some crops that preceded sugarcane had a high effect on sugar yield; this was observed mainly in the first harvest in areas where sunflower, peanuts, velvet beans and sunn hemp were previously cultivated. The 3-year average data showed a sugar yield increase, in the best treatment, of 3 t ha–1 in relation to the control.

The peanut IAC-Caiapó, sunflower and velvet bean were the leguminous crops that resulted in the greater percentage of AM fungus. The lowest population of *Pratylenchus* spp. was

After five harvests, sunn hemp crotalaria was the leguminous crop that induced the greatest

, Raquel Castellucci Caruso Sachs

*Universidade Estadual de Campinas, Departamento de Odontologia SocialPiracicaba (SP) Brazil*

*Faculdade de Zootecnia e Engenharia de Alimentos (FZEA/USP), Pirassununga, (SP), Brazil*

*Centro de Energia Nuclear na Agricultura (CENA/USP), Piracicaba (SP), Brazil*

found in the treatments with peanut IAC-Tatu and IAC-Caiapó.

sugarcane yield, with 30% increase in cane yield and 35% in sugar yield.

the 0-40 cm soil layer with both N sources.

**Author details**

Edmilson José Ambrosano\*

Fábio Luis Ferreira Dias

Gláucia Maria Bovi Ambrosano

Eliana Aparecida Schammas

Heitor Cantarella

Fabrício Rossi

Rozario Azcón

Corresponding Author

 \*

and Juliana Rolim Salomé Teramoto

*APTA, Pólo Regional Centro Sul, Piracicaba (SP), Brazil*

*APTA, Pólo Regional Centro Sul, Piracicaba (SP), Brazil Instituto Agronômico (IAC), Campinas (SP), Brazil*

*Instituto Agronômico (IAC), Campinas (SP), Brazil*

*Instituto de Zootecnia, Nova Odessa (SP), Brazil*

*Estação Experimental de Zaidin, Granada, Espain*

Paulo Cesar Ocheuze Trivelin and Takashi Muraoka

**Figure 3.** Number of nematodes of the genus *Pratylenchus spp*. by 10 grams of roots of sugar sugarcane cultivation influenced by the previous legume species. Adapted from [16].

### **6. Conclusions**

Crop rotation can positively affect yield and increase profit, contributed to raise the net income. This was true both for the green manures (sunn hemp and velvet bean), as for the grain crops (soybean, sunflower and mung bean). Peanuts caused an increase in the sugarcane stalk yields relative to the control, especially in the first harvest, but the high cost of production of this grain somewhat cancelled out the benefit of this rotation.

However, in many sugarcane regions in São Paulo State (Brazil) peanuts are extensively grown in rotation with sugarcane, probably because in those sites yields are higher and the cost of production, lower. Mung beans are a niche crop. Although it provided a relatively high net return in the present study, the risks may be high due to the market restrictions and price fluctuations.

The biomass of green manure induced a complete N substitution in sugarcane and can cause positively affect yield and increase Ca and Mg contents, sum of bases, pH, and base saturation, and decreasing potential acidity and increase profit.

The combination of inorganic fertilizer and green manure resulted in higher sugarcane yields than either N source separately. The recovery of N from ammonium sulfate was higher in the first year whereas in the green manure presented a longer residual effect and resulted in higher yields of cane in the second cycle. The recovery of 15N- labeled fertilizers by two successive sugarcane crops summed up 19 to 21% of the N applied as sunn hemp and 46 to 49% of the N applied as ammonium sulfate. Very little inorganic N was present in the 0-40 cm soil layer with both N sources.

The sugar content of sugarcane stalks is important because the raw material remuneration takes into account this parameter in Brazil. Some crops that preceded sugarcane had a high effect on sugar yield; this was observed mainly in the first harvest in areas where sunflower, peanuts, velvet beans and sunn hemp were previously cultivated. The 3-year average data showed a sugar yield increase, in the best treatment, of 3 t ha–1 in relation to the control.

The peanut IAC-Caiapó, sunflower and velvet bean were the leguminous crops that resulted in the greater percentage of AM fungus. The lowest population of *Pratylenchus* spp. was found in the treatments with peanut IAC-Tatu and IAC-Caiapó.

After five harvests, sunn hemp crotalaria was the leguminous crop that induced the greatest sugarcane yield, with 30% increase in cane yield and 35% in sugar yield.

### **Author details**

26 Biomass Now – Cultivation and Utilization

**6. Conclusions**

price fluctuations.

Thet peanut IAC-Caiapó showed a minimum of 10 nematodes per 10 g of roots and a maximum of 470, while on the control this variation was from 80 to 2,510, indicating the

**Figure 3.** Number of nematodes of the genus *Pratylenchus spp*. by 10 grams of roots of sugar sugarcane

Crop rotation can positively affect yield and increase profit, contributed to raise the net income. This was true both for the green manures (sunn hemp and velvet bean), as for the grain crops (soybean, sunflower and mung bean). Peanuts caused an increase in the sugarcane stalk yields relative to the control, especially in the first harvest, but the high cost

However, in many sugarcane regions in São Paulo State (Brazil) peanuts are extensively grown in rotation with sugarcane, probably because in those sites yields are higher and the cost of production, lower. Mung beans are a niche crop. Although it provided a relatively high net return in the present study, the risks may be high due to the market restrictions and

The biomass of green manure induced a complete N substitution in sugarcane and can cause positively affect yield and increase Ca and Mg contents, sum of bases, pH, and base

The combination of inorganic fertilizer and green manure resulted in higher sugarcane yields than either N source separately. The recovery of N from ammonium sulfate was higher in the first year whereas in the green manure presented a longer residual effect and

of production of this grain somewhat cancelled out the benefit of this rotation.

cultivation influenced by the previous legume species. Adapted from [16].

saturation, and decreasing potential acidity and increase profit.

smaller presence of the nematode in treatments with peanut IAC-Caiapó (Figure 3).

Edmilson José Ambrosano\* , Raquel Castellucci Caruso Sachs and Juliana Rolim Salomé Teramoto *APTA, Pólo Regional Centro Sul, Piracicaba (SP), Brazil*

Fábio Luis Ferreira Dias *APTA, Pólo Regional Centro Sul, Piracicaba (SP), Brazil Instituto Agronômico (IAC), Campinas (SP), Brazil*

Heitor Cantarella *Instituto Agronômico (IAC), Campinas (SP), Brazil*

Gláucia Maria Bovi Ambrosano *Universidade Estadual de Campinas, Departamento de Odontologia SocialPiracicaba (SP) Brazil*

Eliana Aparecida Schammas *Instituto de Zootecnia, Nova Odessa (SP), Brazil*

Fabrício Rossi *Faculdade de Zootecnia e Engenharia de Alimentos (FZEA/USP), Pirassununga, (SP), Brazil*

Paulo Cesar Ocheuze Trivelin and Takashi Muraoka *Centro de Energia Nuclear na Agricultura (CENA/USP), Piracicaba (SP), Brazil*

Rozario Azcón *Estação Experimental de Zaidin, Granada, Espain*

<sup>\*</sup> Corresponding Author

### **Acknowledgement**

To the technical research support of Gilberto Farias, Benedito Mota, and Maria Aparecida C. de Godoy. To FAPESP and CNPq for the grants. Piraí seeds for green manure and cover crops for the support.

Crop Rotation Biomass and Effects on Sugarcane Yield in Brazil 29

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**Chapter 2**

© 2013 Yu et al., licensee InTech. This is an open access chapter distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

distribution, and reproduction in any medium, provided the original work is properly cited.

© 2013 Yu et al., licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

**Generation and Utilization of Microbial Biomass**

In moving towards sustainable manufacturing with reduced carbon footprint, bio-based fuels, chemicals and materials produced from renewable resources have attracted great interest. Microbial cells, in working with other chemical and enzymatic catalysts, are often used in the conversion of feedstocks to desired products, involving different species of bacteria, yeast, filamentous fungi, and microalgae. A substantial amount of microbial biomass is generated in the industrial fermentations and often discarded as a waste. Because of a high cost associated with growth and disposal of the cell mass, reusing the microbial biomass may be an attractive alternative to waste disposal. In contrast to the biomass as energy storage (e.g. starch and oil) or plant structure (e.g. cellulose and hemi-cellulose), microbial biomass is biologically active, consisting primarily of proteins (10-60 wt%), nucleic acid (1-30 wt%), and lipids (1-15%) [1]. Few cases of reusing microbial biomass exist in

Poly(3-hydroxybutyrate) (PHB) is a representative polyhydroxyalkanoate (PHA) that is formed by many bacterial species as carbon and energy reserve [2,3]. Although the biopolyesters made from renewable feedstocks have the potential of replacing petroleumbased thermoplastics in many environmentally friendly applications, they are not widely accepted in the markets because of the high production cost [4]. Extensive research has been conducted to use cheap feedstocks [5,6], develop high cell density fermentation technology for high PHA productivity [7,8], and improve microbial strains that exhibit good performance under high osmotic pressure and environmental stress [9-11]. One major cost factor of PHA production is the recovery and purification of biopolyester for desired purity and material properties [4, 12]. Depending on strains and culture conditions, biopolyester may account for 50-80 wt% of cell mass [13]. They are stored in microbial cells as tiny

**Hydrolysates in Recovery and Production of**

**Poly(3-hydroxybutyrate)**

Jian Yu, Michael Porter and Matt Jaremko

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/52940

**1. Introduction**

industrial processes.

## **Generation and Utilization of Microbial Biomass Hydrolysates in Recovery and Production of Poly(3-hydroxybutyrate)**

Jian Yu, Michael Porter and Matt Jaremko

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/52940

### **1. Introduction**

32 Biomass Now – Cultivation and Utilization

96:1443-1448, 2009

Geoderma, 116:235-248, 2003.

labelled green manures Plant Soil. 322:251–262.

[55] Basanta, M.V.; Dourado Neto, D.; Reichardt, K.; Bacchi, O.O.S.; Oliveira, J.C.M. Trivelin, P.C.O. Timm, L.C.; Tominaga, T.T.; Correchel, V.; Cassaro, F.A.M.; Pires, L.F.; Macedo, J.R. Management effects on nitrogen recovery in a sugarcane crop grown in Brazil.

[56] Asagi, N.; Ueno H. 2009. Nitrogen dynamics in paddy soil applied with various 15N-

[57] Ghiberto, P.J.L. Libardi, P.L.; Brito, A.S.; Trivelin, P.C.O. Leaching of nutrients from a sugarcane crop growing on an Ultisol in Brazil. Agricultural Water Management,

[58] Novaretti, W.R.T. Nematóides em cana-de-açúcar e seu controle. Informe

Agropecuário, v.16, p.37-42. 1992. (in Portuguese, with abstract in English).

In moving towards sustainable manufacturing with reduced carbon footprint, bio-based fuels, chemicals and materials produced from renewable resources have attracted great interest. Microbial cells, in working with other chemical and enzymatic catalysts, are often used in the conversion of feedstocks to desired products, involving different species of bacteria, yeast, filamentous fungi, and microalgae. A substantial amount of microbial biomass is generated in the industrial fermentations and often discarded as a waste. Because of a high cost associated with growth and disposal of the cell mass, reusing the microbial biomass may be an attractive alternative to waste disposal. In contrast to the biomass as energy storage (e.g. starch and oil) or plant structure (e.g. cellulose and hemi-cellulose), microbial biomass is biologically active, consisting primarily of proteins (10-60 wt%), nucleic acid (1-30 wt%), and lipids (1-15%) [1]. Few cases of reusing microbial biomass exist in industrial processes.

Poly(3-hydroxybutyrate) (PHB) is a representative polyhydroxyalkanoate (PHA) that is formed by many bacterial species as carbon and energy reserve [2,3]. Although the biopolyesters made from renewable feedstocks have the potential of replacing petroleumbased thermoplastics in many environmentally friendly applications, they are not widely accepted in the markets because of the high production cost [4]. Extensive research has been conducted to use cheap feedstocks [5,6], develop high cell density fermentation technology for high PHA productivity [7,8], and improve microbial strains that exhibit good performance under high osmotic pressure and environmental stress [9-11]. One major cost factor of PHA production is the recovery and purification of biopolyester for desired purity and material properties [4, 12]. Depending on strains and culture conditions, biopolyester may account for 50-80 wt% of cell mass [13]. They are stored in microbial cells as tiny

© 2013 Yu et al., licensee InTech. This is an open access chapter distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 Yu et al., licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

amorphous granules [0.2-0.5 m in diameter] and need a sophisticated treatment to separate them from the residual cell mass [14-16]. Two technologies, based-on solvent extraction or biomass dissolution, are usually adopted in PHA recovery. With solvent extraction, the PHA granules are dissolved in appropriate organic solvents, leaving cells or residual biomass intact [17,18]. With cell mass dissolution, the PHA granules are left intact while the non-PHA cell mass is decomposed and dissolved in aqueous solutions with help of biological and/or chemical agents [19,20]. Following either treatment, PHA and non-PHA cell mass can be separated with conventional solid/liquid separations. Separating biopolyester from the cells would generate a substantial amount of residual microbial biomass, 0.25 to 1 kg dry mass per kg of PHA resin, depending on the initial PHA content. As a mixture of proteins, nucleic acids, lipids, and wall fragments, the residual biomass has no market value and is discarded at an extra disposal cost.

Generation and Utilization of Microbial Biomass

Hydrolysates in Recovery and Production of Poly(3-hydroxybutyrate) 35

hydrolysates, and the rest pre-sterilized water. The glucose mineral solution contained (per liter): 12 g glucose, 2 g NaH2PO4, 2.8 g K2HPO4, 0.5 g MgSO4.7H2O, 1 g (NH4)2SO4 and 1 mL trace solution [21]. The culture solution was shaken in a 500 mL baffled flask at 30 oC and 200 rpm in a rotary incubator for 24 or 48 hours. Cell mass was harvested from 50 mL medium with centrifugation at 5,000 g for 10 min and freeze dried for measurement of cell

A large amount of PHB-containing cell mass for biopolyester recovery was produced with a fed-batch culture in a 3L bench-top bioreactor (BioFlo 110, New Brunswick Scientific Co. NJ). The temperature, pH and dissolved oxygen were controlled at 30 oC, 6.8, and 10% air situation, respectively. A feed solution prepared with a sugar manufacturing byproduct containing about 50% sucrose was introduced into the bioreactor till the cell density reached 130 g/L and PHB concentration 94 g/L (72% PHB of cell mass). The cell mass was harvested with centrifugation and re-suspended in an acidic water (0.2M H2SO4) to make a cell slurry

The biopolyester was recovered and purified by dissolving the non-PHA cell mass (28% w/w) in sequential treatments consisting of acid pretreatment, base treatment, hypochlorite

Acid pretreatment: The cell mass in acidic solution was heated to boil and maintained for one hour under ambient conditions. The pretreated cellular solids were cooled to room temperature and separated from acid solution with centrifugation at 5,000 g for 10 min. The dissolved microbial biomass in the supernatant solution is referred to acid hydrolysates. The

Base treatment: The insoluble solids from the acid pretreatment were re-suspended in an equivalent volume of water to form slurry of about 200 g DM/L. The solution pH was raised to 10 to 11 with 5 M NaOH solution and stirred under ambient conditions for 30 to 60 min. A small amount of surfactants such as sodium dodecylsulfate (SDS, CH3(CH2)11OSO3Na) might be added to a concentration of 5 to 10 g/L. The slurry was then heated to boiling and maintained for 10 min under ambient conditions. After centrifugation, the dissolved biomass in the supernatant solution is referred to base hydrolysates. The sequential acid and base treatments above could also be performed without separation of the acid hydrolysates. In this case, sodium hydroxide was directly added into the acidic cell slurry and the pH was raised to 10-11 for base treatment. After centrifugation, the supernatant solution contained the hydrolysates generated from both acid and base treatments and is refereed to acid-base

Whitening and washing: The insoluble wet pellets from base treatment were re-suspended in a commercially available bleaching solution containing 6% w/w of hypochlorite. The slurry was stirred for 1 to 2 hours under ambient conditions. The white PHB pellets were recovered with centrifugation. A small amount of biomass was dissolved and mineralized in

mass concentration and PHB content.

whitening, washing and drying [22].

**2.2. PHB recovery**

hydrolysates.

of 278 g dry mass/L. The slurry was reserved for later use.

insoluble wet pellets were subjected to next base treatment.

According to the microbial structure and cellular composition [1], the residual microbial biomass is actually consisting of true biological compounds formed during cell growth while PHA is just a carbon storage material. In a conventional PHA fermentation, sufficient substrates and nutrients (C, N, P, minerals and some organic growth factors) are supplied to grow enough cell mass that in turn or simultaneously synthesize biopolyester from carbon substrates. A large portion of organic carbons and nutrients are therefore consumed to generate new cell mass that is going to be discarded as a solid waste after polymer recovery. Ideally, the residual biomass should be reused by microbial cells to generate new cell mass and/or PHA polymers. This would not only reduce the cost of waste treatment and disposal, but also save the cost of nutrients in PHA fermentation. The bacterial cells, however, cannot assimilate their cell mass because they lack appropriate enzymes to break down various biological macromolecules and their complex structure such as cellular walls and membranes [19]. If the cells or mutants from genetic engineering could easily assimilate their structural components, they might not be suitable to industrial PHA production because the cells would undergo autolysis under high environmental stress. It is highly possible, however, to make the residual biomass reusable during PHA recovery in which the non-PHA cell mass is decomposed and hydrolyzed in aqueous solutions [20]. Integration of PHA recovery with reusing of residual biomass in microbial PHA fermentation is a novel and challenging technology. This work shows the generation of biomass hydrolysates in a downstream PHB recovery and the beneficial utilization of the hydrolysates in cell growth and PHB formation.

### **2. Materials and methods**

### **2.1. Microbial cultures**

A laboratory strain of *Ralstonia eutropha* maintained on agar slant was used in this work. The agar medium contained (per liter): 5 g yeast extract, 5 g peptone, 2.5 g meat extract and 15 g agar. A 16S RNA analysis indicates that the strain has 100% alignment with *Ralstonia eutropha* H16. The cells were cultivated in 200 mL mineral medium containing: 160 mL glucose mineral solution, 8 mL inoculum, 5-20 mL solutions of residual microbial biomass hydrolysates, and the rest pre-sterilized water. The glucose mineral solution contained (per liter): 12 g glucose, 2 g NaH2PO4, 2.8 g K2HPO4, 0.5 g MgSO4.7H2O, 1 g (NH4)2SO4 and 1 mL trace solution [21]. The culture solution was shaken in a 500 mL baffled flask at 30 oC and 200 rpm in a rotary incubator for 24 or 48 hours. Cell mass was harvested from 50 mL medium with centrifugation at 5,000 g for 10 min and freeze dried for measurement of cell mass concentration and PHB content.

A large amount of PHB-containing cell mass for biopolyester recovery was produced with a fed-batch culture in a 3L bench-top bioreactor (BioFlo 110, New Brunswick Scientific Co. NJ). The temperature, pH and dissolved oxygen were controlled at 30 oC, 6.8, and 10% air situation, respectively. A feed solution prepared with a sugar manufacturing byproduct containing about 50% sucrose was introduced into the bioreactor till the cell density reached 130 g/L and PHB concentration 94 g/L (72% PHB of cell mass). The cell mass was harvested with centrifugation and re-suspended in an acidic water (0.2M H2SO4) to make a cell slurry of 278 g dry mass/L. The slurry was reserved for later use.

### **2.2. PHB recovery**

34 Biomass Now – Cultivation and Utilization

no market value and is discarded at an extra disposal cost.

hydrolysates in cell growth and PHB formation.

**2. Materials and methods**

**2.1. Microbial cultures**

amorphous granules [0.2-0.5 m in diameter] and need a sophisticated treatment to separate them from the residual cell mass [14-16]. Two technologies, based-on solvent extraction or biomass dissolution, are usually adopted in PHA recovery. With solvent extraction, the PHA granules are dissolved in appropriate organic solvents, leaving cells or residual biomass intact [17,18]. With cell mass dissolution, the PHA granules are left intact while the non-PHA cell mass is decomposed and dissolved in aqueous solutions with help of biological and/or chemical agents [19,20]. Following either treatment, PHA and non-PHA cell mass can be separated with conventional solid/liquid separations. Separating biopolyester from the cells would generate a substantial amount of residual microbial biomass, 0.25 to 1 kg dry mass per kg of PHA resin, depending on the initial PHA content. As a mixture of proteins, nucleic acids, lipids, and wall fragments, the residual biomass has

According to the microbial structure and cellular composition [1], the residual microbial biomass is actually consisting of true biological compounds formed during cell growth while PHA is just a carbon storage material. In a conventional PHA fermentation, sufficient substrates and nutrients (C, N, P, minerals and some organic growth factors) are supplied to grow enough cell mass that in turn or simultaneously synthesize biopolyester from carbon substrates. A large portion of organic carbons and nutrients are therefore consumed to generate new cell mass that is going to be discarded as a solid waste after polymer recovery. Ideally, the residual biomass should be reused by microbial cells to generate new cell mass and/or PHA polymers. This would not only reduce the cost of waste treatment and disposal, but also save the cost of nutrients in PHA fermentation. The bacterial cells, however, cannot assimilate their cell mass because they lack appropriate enzymes to break down various biological macromolecules and their complex structure such as cellular walls and membranes [19]. If the cells or mutants from genetic engineering could easily assimilate their structural components, they might not be suitable to industrial PHA production because the cells would undergo autolysis under high environmental stress. It is highly possible, however, to make the residual biomass reusable during PHA recovery in which the non-PHA cell mass is decomposed and hydrolyzed in aqueous solutions [20]. Integration of PHA recovery with reusing of residual biomass in microbial PHA fermentation is a novel and challenging technology. This work shows the generation of biomass hydrolysates in a downstream PHB recovery and the beneficial utilization of the

A laboratory strain of *Ralstonia eutropha* maintained on agar slant was used in this work. The agar medium contained (per liter): 5 g yeast extract, 5 g peptone, 2.5 g meat extract and 15 g agar. A 16S RNA analysis indicates that the strain has 100% alignment with *Ralstonia eutropha* H16. The cells were cultivated in 200 mL mineral medium containing: 160 mL glucose mineral solution, 8 mL inoculum, 5-20 mL solutions of residual microbial biomass The biopolyester was recovered and purified by dissolving the non-PHA cell mass (28% w/w) in sequential treatments consisting of acid pretreatment, base treatment, hypochlorite whitening, washing and drying [22].

Acid pretreatment: The cell mass in acidic solution was heated to boil and maintained for one hour under ambient conditions. The pretreated cellular solids were cooled to room temperature and separated from acid solution with centrifugation at 5,000 g for 10 min. The dissolved microbial biomass in the supernatant solution is referred to acid hydrolysates. The insoluble wet pellets were subjected to next base treatment.

Base treatment: The insoluble solids from the acid pretreatment were re-suspended in an equivalent volume of water to form slurry of about 200 g DM/L. The solution pH was raised to 10 to 11 with 5 M NaOH solution and stirred under ambient conditions for 30 to 60 min. A small amount of surfactants such as sodium dodecylsulfate (SDS, CH3(CH2)11OSO3Na) might be added to a concentration of 5 to 10 g/L. The slurry was then heated to boiling and maintained for 10 min under ambient conditions. After centrifugation, the dissolved biomass in the supernatant solution is referred to base hydrolysates. The sequential acid and base treatments above could also be performed without separation of the acid hydrolysates. In this case, sodium hydroxide was directly added into the acidic cell slurry and the pH was raised to 10-11 for base treatment. After centrifugation, the supernatant solution contained the hydrolysates generated from both acid and base treatments and is refereed to acid-base hydrolysates.

Whitening and washing: The insoluble wet pellets from base treatment were re-suspended in a commercially available bleaching solution containing 6% w/w of hypochlorite. The slurry was stirred for 1 to 2 hours under ambient conditions. The white PHB pellets were recovered with centrifugation. A small amount of biomass was dissolved and mineralized in the bleaching solution because of chemical oxidation, which is not considered for reuse in this work. The wet PHA pellets were washed two times with water and dried in oven. The final PHB product is a white powder.

Generation and Utilization of Microbial Biomass

Hydrolysates in Recovery and Production of Poly(3-hydroxybutyrate) 37

**Figure 1.** Sequential treatment of PHB-containing biomass (100 g dry mass) and generation of biomass

**Figure 2.** Effect of temperature and time on release of proteins from microbial cells in acidic solution.

During the acid pretreatment, the original amorphous PHB granules became partially crystallized (data not shown here), which improved the granule's resistance to abiotic degradation in the following treatments [20]. The biomass hydrolysates (13.4 g dry mass) dissolved in supernatant solution was discharged as acid hydrolysates. The residual PHBcontaining biomass was further subjected to a base treatment by raising the slurry pH to 10.5 with a 10M NaOH solution. About 15.4 g dry mass was dissolved in the supernatant solution and discharged as base hydrolysates. After a small amount of residual biomass (2 g dry mass) was removed via oxidation with hypochlorite, the final PHB powder (69.2 g dry mass) contained 96.4 wt% PHB. The overall PHB recovery yield was 92.6%, or 7-8% PHB

hydrolysates. Surfactant SDS is optional for high PHB purity.

### **2.3. Chemical and material analysis**

The dissolution of non-PHA cell mass was monitored by measuring the characteristic absorption of amino acid residues at 280 nm with a UV/VIS spectrophotometer (Beckman Coulter DU530, Fullerton, CA). The concentration of proteins that can be stained in Bradford assay was measured with the spectrophotometer after protein-dye binding [23]. The content of PHB in original cell mass, in sequential treatments, and final product were determined via methanolysis of the biopolyester in methanol (3 wt% sulfuric acid) at 100 oC for 8-10 hours [24]. The 3-hydroxybutyric methyl ester was hydrolyzed into 3-hydroxybutyric acid when the solution pH was raised to 11 with a 10N NaOH solution. The liquid samples were analyzed using an HPLC equipped with a UV detector (Shimadzu, Japan) and an organic acid column (OA-1000, Alltech, Deerfield, IL). The column was maintained at 65 oC and eluted with a water-sulfuric acid solution (pH 2) at 0.8 mL/min. The monomeric acid and crotonic acid, a trace byproduct formed in methanolysis, were detected at 210 nm. For data quality control, the biopolyester was also extracted from the freeze-dried cell mass in hot chloroform followed by precipitation with methanol [21]. The PHB content was calculated from the purified PHB and compared with the results of HPLC analysis.

The purified PHB and non-PHB cell mass were examined with a Nicolet Avatar 370 FTIR spectrometer (Thermo Electron Co., Madison, WI). The solids were pressed on a germanium crystal window of micro-horizontal attenuated total reflectance (ATR) for measurement of single-reflection and absorption of infrared radiation by the specimens. The thermal properties of PHB powder were examined with a differential scanning calorimeter (DSC). A Modulated 2920 instrument (TA Instruments, New Castle, DE) equipped with a refrigerated cooling system was run in heat-cool-heat mode at a rate of 5 oC/min under nitrogen. The selected temperature range was 30oC – 210 oC with sample weights of 4.5 – 5.5 mgs. Images of cell and PHB granules were obtained with an energyfiltering transmission electron microscopy (120 kV LEO 912, Carl Zeiss SMT Inc. MA). The instrument has an in-column electron energy loss spectrometer, allowing analysis of light element in thin sections.

### **3. Results and discussion**

### **3.1. Sequential treatment for PHB recovery**

Figure 1 elucidates the sequential treatment of PHB-containing cells in a process of PHB recovery and purification. Starting with 100 dry cell mass, the cells in a slurry of 278 g DM/L were first treated in an acidic solution (0.2M H2SO4). A substantial amount of microbial proteins was released from the damaged cells, depending on temperature and time as shown in Figure 2.

Generation and Utilization of Microbial Biomass

Hydrolysates in Recovery and Production of Poly(3-hydroxybutyrate) 37

36 Biomass Now – Cultivation and Utilization

element in thin sections.

shown in Figure 2.

**3. Results and discussion**

**3.1. Sequential treatment for PHB recovery**

final PHB product is a white powder.

**2.3. Chemical and material analysis**

the bleaching solution because of chemical oxidation, which is not considered for reuse in this work. The wet PHA pellets were washed two times with water and dried in oven. The

The dissolution of non-PHA cell mass was monitored by measuring the characteristic absorption of amino acid residues at 280 nm with a UV/VIS spectrophotometer (Beckman Coulter DU530, Fullerton, CA). The concentration of proteins that can be stained in Bradford assay was measured with the spectrophotometer after protein-dye binding [23]. The content of PHB in original cell mass, in sequential treatments, and final product were determined via methanolysis of the biopolyester in methanol (3 wt% sulfuric acid) at 100 oC for 8-10 hours [24]. The 3-hydroxybutyric methyl ester was hydrolyzed into 3-hydroxybutyric acid when the solution pH was raised to 11 with a 10N NaOH solution. The liquid samples were analyzed using an HPLC equipped with a UV detector (Shimadzu, Japan) and an organic acid column (OA-1000, Alltech, Deerfield, IL). The column was maintained at 65 oC and eluted with a water-sulfuric acid solution (pH 2) at 0.8 mL/min. The monomeric acid and crotonic acid, a trace byproduct formed in methanolysis, were detected at 210 nm. For data quality control, the biopolyester was also extracted from the freeze-dried cell mass in hot chloroform followed by precipitation with methanol [21]. The PHB content was calculated

The purified PHB and non-PHB cell mass were examined with a Nicolet Avatar 370 FTIR spectrometer (Thermo Electron Co., Madison, WI). The solids were pressed on a germanium crystal window of micro-horizontal attenuated total reflectance (ATR) for measurement of single-reflection and absorption of infrared radiation by the specimens. The thermal properties of PHB powder were examined with a differential scanning calorimeter (DSC). A Modulated 2920 instrument (TA Instruments, New Castle, DE) equipped with a refrigerated cooling system was run in heat-cool-heat mode at a rate of 5 oC/min under nitrogen. The selected temperature range was 30oC – 210 oC with sample weights of 4.5 – 5.5 mgs. Images of cell and PHB granules were obtained with an energyfiltering transmission electron microscopy (120 kV LEO 912, Carl Zeiss SMT Inc. MA). The instrument has an in-column electron energy loss spectrometer, allowing analysis of light

Figure 1 elucidates the sequential treatment of PHB-containing cells in a process of PHB recovery and purification. Starting with 100 dry cell mass, the cells in a slurry of 278 g DM/L were first treated in an acidic solution (0.2M H2SO4). A substantial amount of microbial proteins was released from the damaged cells, depending on temperature and time as

from the purified PHB and compared with the results of HPLC analysis.

**Figure 1.** Sequential treatment of PHB-containing biomass (100 g dry mass) and generation of biomass hydrolysates. Surfactant SDS is optional for high PHB purity.

**Figure 2.** Effect of temperature and time on release of proteins from microbial cells in acidic solution.

During the acid pretreatment, the original amorphous PHB granules became partially crystallized (data not shown here), which improved the granule's resistance to abiotic degradation in the following treatments [20]. The biomass hydrolysates (13.4 g dry mass) dissolved in supernatant solution was discharged as acid hydrolysates. The residual PHBcontaining biomass was further subjected to a base treatment by raising the slurry pH to 10.5 with a 10M NaOH solution. About 15.4 g dry mass was dissolved in the supernatant solution and discharged as base hydrolysates. After a small amount of residual biomass (2 g dry mass) was removed via oxidation with hypochlorite, the final PHB powder (69.2 g dry mass) contained 96.4 wt% PHB. The overall PHB recovery yield was 92.6%, or 7-8% PHB

was lost in repeated hydrolysis and solid/liquid separations. When a small amount of surfactant such as sodium dodecylsulfate (SDS) was added in the base treatment, the PHB purity of the final biopolyester resin could be increased to 99.4%.

Generation and Utilization of Microbial Biomass

Hydrolysates in Recovery and Production of Poly(3-hydroxybutyrate) 39

Figure 4 compares the FTIR spectra of the purified PHB granules and the original oven-dried PHB-containing cell mass. The peaks of amide I band at ~1650 cm-1 and amide II band at ~1540 cm-1 (N-H bend) are characteristic infrared radiation absorption of the proteins in cell mass [25,26]. They disappeared in the spectrum of purified PHB granules. This was confirmed with a pure PHB prepared with solvent extraction (the spectrum not shown here). It was also noticed that the native amorphous PHB granules became crystallized during the process of purification, which will be further discussed. This structural change in PHB matrix was also reflected in the

absorption of infrared radiation at wave numbers of 1180, 1210, and 1280 nm-1 [27].

**Figure 4.** FTIR spectra of purified PHB granules and PHB-containing oven-dried cell mass (ODCM).

The purified PHB granules were subjected to repeated heating and cooling in a differential scanning calorimeter (DSC), and the results are presented in Figure 5. In the first heating (solid blue line), two melting peaks were observed, around 156 oC and 167 oC, respectively, indicating that the PHB powder had two types of crystalline structures. The melted polymer was re-crystallized again during the first cooling (dotted blue line), starting at 100 oC and ending at 80 oC. When the crystallized PHB was subjected to the second heating (solid black line), it was noticed that the relatively small melting peak at 156 oC (or crystalline structure) observed in the first heating disappeared. Only one melting peak (crystalline structure) was observed, starting at 160 oC and ending at 181 oC with a peak around 174 oC. This phenomenon indicates that the first melting peak in the first heating represents a type of crystalline structure that could not be formed at a cooling rate of 5 oC/min. The whole endothermic event in the second heating absorbed 87J/g PHB. Based on a theoretical melting

**3.2. Purified PHB versus original cell mass**

**Figure 3.** Transmission electron microscope images: microbial cells containing native PHB granules (top left), cells with damaged walls in acid pretreatment (top right), PHB granules with attached residual cell mass (bottom left), and purified PHB granules (bottom right)

Figure 3 is the electronic microscopy images of the original cells with PHB inclusion bodies, the cells with damaged porous cell walls in acid pretreatment, the PHB granules with residual cellular mass in base treatment, and the purified PHB granules after whitening and washing. It is interesting to see that the cell walls became porous in the acid pretreatment, which allowed release of proteins and other biological components in cytoplasm. The original cell structure, however, was maintained to keep the PHB granules within the damaged cells. After base treatment, the cell walls were almost completely decomposed, and a small amount of residual cell mass, probably some hydrophobic cellular components, was attached to PHB granules. After whitening and washing, the non-PHA cell mass was removed to give purified PHB granules.

### **3.2. Purified PHB versus original cell mass**

38 Biomass Now – Cultivation and Utilization

was lost in repeated hydrolysis and solid/liquid separations. When a small amount of surfactant such as sodium dodecylsulfate (SDS) was added in the base treatment, the PHB

**Figure 3.** Transmission electron microscope images: microbial cells containing native PHB granules (top left), cells with damaged walls in acid pretreatment (top right), PHB granules with attached

Figure 3 is the electronic microscopy images of the original cells with PHB inclusion bodies, the cells with damaged porous cell walls in acid pretreatment, the PHB granules with residual cellular mass in base treatment, and the purified PHB granules after whitening and washing. It is interesting to see that the cell walls became porous in the acid pretreatment, which allowed release of proteins and other biological components in cytoplasm. The original cell structure, however, was maintained to keep the PHB granules within the damaged cells. After base treatment, the cell walls were almost completely decomposed, and a small amount of residual cell mass, probably some hydrophobic cellular components, was attached to PHB granules. After whitening and washing, the non-PHA cell mass was

residual cell mass (bottom left), and purified PHB granules (bottom right)

removed to give purified PHB granules.

purity of the final biopolyester resin could be increased to 99.4%.

Figure 4 compares the FTIR spectra of the purified PHB granules and the original oven-dried PHB-containing cell mass. The peaks of amide I band at ~1650 cm-1 and amide II band at ~1540 cm-1 (N-H bend) are characteristic infrared radiation absorption of the proteins in cell mass [25,26]. They disappeared in the spectrum of purified PHB granules. This was confirmed with a pure PHB prepared with solvent extraction (the spectrum not shown here). It was also noticed that the native amorphous PHB granules became crystallized during the process of purification, which will be further discussed. This structural change in PHB matrix was also reflected in the absorption of infrared radiation at wave numbers of 1180, 1210, and 1280 nm-1 [27].

**Figure 4.** FTIR spectra of purified PHB granules and PHB-containing oven-dried cell mass (ODCM).

The purified PHB granules were subjected to repeated heating and cooling in a differential scanning calorimeter (DSC), and the results are presented in Figure 5. In the first heating (solid blue line), two melting peaks were observed, around 156 oC and 167 oC, respectively, indicating that the PHB powder had two types of crystalline structures. The melted polymer was re-crystallized again during the first cooling (dotted blue line), starting at 100 oC and ending at 80 oC. When the crystallized PHB was subjected to the second heating (solid black line), it was noticed that the relatively small melting peak at 156 oC (or crystalline structure) observed in the first heating disappeared. Only one melting peak (crystalline structure) was observed, starting at 160 oC and ending at 181 oC with a peak around 174 oC. This phenomenon indicates that the first melting peak in the first heating represents a type of crystalline structure that could not be formed at a cooling rate of 5 oC/min. The whole endothermic event in the second heating absorbed 87J/g PHB. Based on a theoretical melting enthalpy of 100% crystalline PHB (146 J/g) [28], it can be estimated that about 60% of PHB matrix was crystallized during the first cooling (Eq. 1).

$$X\_{\mathcal{C}} = \frac{\Delta H\_m}{\Delta H\_l} = \frac{87}{146} \approx 60\% \tag{1}$$

Generation and Utilization of Microbial Biomass

Hydrolysates in Recovery and Production of Poly(3-hydroxybutyrate) 41

oxidation with hypochlorite, a strong oxidation agent. The acid hydrolysates are primarily the cytoplasm proteins released from the damaged cells (Figures 2 and 3). The released biological macromolecules were subjected to further hydrolysis in the thermal acidic solution. The acid hydrolysates solution had a clean brownish color and contained 30-45 g/ of soluble biomass, depending on the density of cell slurry and treatment conditions. The base hydrolysates solution with a dark color contained the hydrolysis products of hydrophobic cell components including lipids and membrane proteins. After centrifugation, the concentration of soluble biomass in the supernatant solution was 30 to 50 g/L. The sequential treatments disrupted and dissolved the structural components so that they could be removed from PHB granules. Equally important, the biomass and biological macromolecules were decomposed into small molecule hydrolysates such as amino acids and organic acids. These hydrolysates could become appropriate substrates that can be

assimilated by microbial cells in microbial PHB production.

**Figure 6.** FTIR spectra of acid-base biomass hydrolysates (black line, cell debris) and cellular

In addition to the two types of biomass hydrolysates described above, a mixed hydrolysates of residual biomass was generated when the acid pretreatment and base treatment were performed sequentially without solid/liquid separation. It eliminated one operation of solid/liquid separation, but the acid hydrolysates (primarily proteins) were subjected to additional hydrolysis in the base solution. Figure 6 shows the FTIR spectrum of an acid-base hydrolysates. As observed in the IR spectrum of the original cell mass in Figure 4, a major component of the hydrolysates was the amino acids or proteins with infrared radiation absorbance at 1500 to 1700 cm-1 [25]. Another major component in the acid/base hydrolysates had the IR absorbance at 1000 to 1200 cm-1, which was attributed to cell lipids

components extracted with acetone (blue line)

Where Xc is the PHB crystallinity, Hm and Ht are the melting enthalpies of PHB powder and a theoretical PHB crystal [28].

**Figure 5.** Differential scanning calorimetry (DSC) measurement of purified PHB granules in repeated heating and cooling: the first heating (solid blue line) followed by the first cooling (dotted blue line) and the second heating (solid black line) following by the second cooling (dotted black line).

In contrast to the thermal plastic behavior of PHB powders, the oven-dried cell mass containing 73% PHB could not be melted till carbonization. This fact reveals a complicated interaction between the biopolyester and the residual biomass. It also shows that a composite of 73% PHB and 27% cellular mass is not a thermoplastic material, but a rigid composite. The role of cellular mass in the PHB composite is not clear yet.

### **3.3. Cell debris solutions**

As shown in Figure 1, about 94 wt% of residual microbial biomass was decomposed and hydrolyzed in the acid pretreatment (~44% biomass) and base treatment (~50% biomass). The remaining small amount (~6 wt %) of residual cell mass is most likely mineralized via oxidation with hypochlorite, a strong oxidation agent. The acid hydrolysates are primarily the cytoplasm proteins released from the damaged cells (Figures 2 and 3). The released biological macromolecules were subjected to further hydrolysis in the thermal acidic solution. The acid hydrolysates solution had a clean brownish color and contained 30-45 g/ of soluble biomass, depending on the density of cell slurry and treatment conditions. The base hydrolysates solution with a dark color contained the hydrolysis products of hydrophobic cell components including lipids and membrane proteins. After centrifugation, the concentration of soluble biomass in the supernatant solution was 30 to 50 g/L. The sequential treatments disrupted and dissolved the structural components so that they could be removed from PHB granules. Equally important, the biomass and biological macromolecules were decomposed into small molecule hydrolysates such as amino acids and organic acids. These hydrolysates could become appropriate substrates that can be assimilated by microbial cells in microbial PHB production.

40 Biomass Now – Cultivation and Utilization

and a theoretical PHB crystal [28].

**3.3. Cell debris solutions**

matrix was crystallized during the first cooling (Eq. 1).

enthalpy of 100% crystalline PHB (146 J/g) [28], it can be estimated that about 60% of PHB

<sup>=</sup> <sup>87</sup> 146

Where Xc is the PHB crystallinity, Hm and Ht are the melting enthalpies of PHB powder

**Figure 5.** Differential scanning calorimetry (DSC) measurement of purified PHB granules in repeated heating and cooling: the first heating (solid blue line) followed by the first cooling (dotted blue line) and

In contrast to the thermal plastic behavior of PHB powders, the oven-dried cell mass containing 73% PHB could not be melted till carbonization. This fact reveals a complicated interaction between the biopolyester and the residual biomass. It also shows that a composite of 73% PHB and 27% cellular mass is not a thermoplastic material, but a rigid

As shown in Figure 1, about 94 wt% of residual microbial biomass was decomposed and hydrolyzed in the acid pretreatment (~44% biomass) and base treatment (~50% biomass). The remaining small amount (~6 wt %) of residual cell mass is most likely mineralized via

the second heating (solid black line) following by the second cooling (dotted black line).

composite. The role of cellular mass in the PHB composite is not clear yet.

 ≈ 60% (1)

�� <sup>=</sup> ��� ���

**Figure 6.** FTIR spectra of acid-base biomass hydrolysates (black line, cell debris) and cellular components extracted with acetone (blue line)

In addition to the two types of biomass hydrolysates described above, a mixed hydrolysates of residual biomass was generated when the acid pretreatment and base treatment were performed sequentially without solid/liquid separation. It eliminated one operation of solid/liquid separation, but the acid hydrolysates (primarily proteins) were subjected to additional hydrolysis in the base solution. Figure 6 shows the FTIR spectrum of an acid-base hydrolysates. As observed in the IR spectrum of the original cell mass in Figure 4, a major component of the hydrolysates was the amino acids or proteins with infrared radiation absorbance at 1500 to 1700 cm-1 [25]. Another major component in the acid/base hydrolysates had the IR absorbance at 1000 to 1200 cm-1, which was attributed to cell lipids

and/or similar compounds. This was confirmed with the spectrum of cell mass extract in acetone. Acetone is a common solvent used to remove hydrophobic lipids, steroids and pigments from PHB-containing cell mass [17]. It does not dissolve and extract PHB and proteins. Based on the observations above, it was concluded that the major cellular components in the acid hydrolysates were derived from cytoplasm proteins and in the base hydrolysates from cell walls, lipids and membrane proteins. In the acid-base hydrolysates, the products were derived from both groups, i.e. amino acids or peptides derived from proteins, and lipids derived from cell walls and membranes. It should be pointed out that the composition of acid-base hydrolysates is not a simple mixture of acid- and basehydrolysates because the acid hydrolysates were further hydrolyzed in base treatment.

Generation and Utilization of Microbial Biomass

Hydrolysates in Recovery and Production of Poly(3-hydroxybutyrate) 43

(bottom) in reuse of the residual biomass for PHB production.

**Figure 7.** Effect of acid hydrolysates to glucose loading ratio on cell growth (top) and PHB formation

in Table 1, the concentrations of both cell mass and PHA content, after 48 hours cultivation, were substantially higher than those of the control. The overall cell growth yield (Yx/s) and PHA formation yield (Yp/s) are calculated from the amounts of cell mass and PHA formed in 48 hours based on the initial concentration of glucose. The relative yields (Yx' and Yp') based on the controls were increased by 100 – 300%. More interestingly, the inhibitory effect of acid biomass hydrolysates was not observed in the use of acid-base hydrolysates, and the load of cell debris to glucose can be increased to about 39 wt%. Most likely, the unknown inhibitors in acid hydrolysates were further decomposed into less inhibitory hydrolysates in the base treatment. Since the acid biomass hydrolysates contained primarily the cytoplasm proteins released from the damaged cells, the soluble proteins might adversely affect cell growth at high concentrations. After being hydrolyzed in base solution into peptides and amino acids,

the small molecule hydrolysates become less inhibitory and more usable to the cells.

Because of the hydrophobic properties of PHB granules, the residual hydrophobic impurities of cell mass might be attached to the granules and difficult to remove by washing with water. A surfactant such as SDS in the base treatment can remove most of the impurities to a high PHB purity (>99 % w/w). A large portion of the surfactant, however, may be left in the hydrolysates solution and may have an adverse effect on the reuse of the hydrolysates in PHB production.

### **3.4. Utilization of acid hydrolysates in PHB biosynthesis**

An acid hydrolysates solution containing 38 g/L of soluble solids was added into a glucose medium to give a predetermined percentage of residual biomass to glucose at 0, 10, 20 and 25% of sugar, respectively. The initial glucose concentration was controlled at a constant level of 9.6 g/L. The flask cultures of no biomass hydrolysates were run in parallel as controls. As shown in Figure 7, the acid biomass hydrolysates were beneficial to both cell growth and PHA formation. Because the residual biomass might also contain some insoluble solids and PHB granules lost in PHB recovery, both cell density and PHB concentration were compared at 24 hours and 48 hours to show the net gains. The benefits of biomass hydrolysates were statistically significant based on the deviations of duplicates.

The acid hydrolysates might have two positive effects on microbial PHA formation. First, the hydrolysates promoted cell activity on glucose utilization, giving higher cell densities than the controls in the first 24 hours. This nutritional effect was similar to those of organic nutrients such as yeast extract and peptone, which are widely used in microbial cultures to provide nutrients and growth factors to the cells. A fast cell growth can reduce the cultivation time, resulting in a high PHB productivity. Second, the biomass hydrolysates might also be used as an extra carbon source to generate more cell mass than the controls in 48 hours. This carbon source effect, however, might play a minor role because the cell density did not increase with cell debris load. In fact, too much acid hydrolysates deteriorated the gains as shown in Figure 7. The reason is not clear yet. A load of acid biomass hydrolysates to glucose from 10 to 20 wt% seems appropriate for both cell growth and PHB formation.

### **3.5. Utilization of acid-base biomass hydrolysates**

An acid-base biomass hydrolysates solution containing 48.5 g/L of soluble solids was added into a glucose medium at predetermined percentage of cell debris to glucose from 0 to 40%. The glucose medium without biomass hydrolysates was run in parallel as controls. As shown

Generation and Utilization of Microbial Biomass Hydrolysates in Recovery and Production of Poly(3-hydroxybutyrate) 43

42 Biomass Now – Cultivation and Utilization

and/or similar compounds. This was confirmed with the spectrum of cell mass extract in acetone. Acetone is a common solvent used to remove hydrophobic lipids, steroids and pigments from PHB-containing cell mass [17]. It does not dissolve and extract PHB and proteins. Based on the observations above, it was concluded that the major cellular components in the acid hydrolysates were derived from cytoplasm proteins and in the base hydrolysates from cell walls, lipids and membrane proteins. In the acid-base hydrolysates, the products were derived from both groups, i.e. amino acids or peptides derived from proteins, and lipids derived from cell walls and membranes. It should be pointed out that the composition of acid-base hydrolysates is not a simple mixture of acid- and basehydrolysates because the acid hydrolysates were further hydrolyzed in base treatment.

Because of the hydrophobic properties of PHB granules, the residual hydrophobic impurities of cell mass might be attached to the granules and difficult to remove by washing with water. A surfactant such as SDS in the base treatment can remove most of the impurities to a high PHB purity (>99 % w/w). A large portion of the surfactant, however, may be left in the hydrolysates solution and may have an adverse effect on the reuse of the hydrolysates in PHB production.

An acid hydrolysates solution containing 38 g/L of soluble solids was added into a glucose medium to give a predetermined percentage of residual biomass to glucose at 0, 10, 20 and 25% of sugar, respectively. The initial glucose concentration was controlled at a constant level of 9.6 g/L. The flask cultures of no biomass hydrolysates were run in parallel as controls. As shown in Figure 7, the acid biomass hydrolysates were beneficial to both cell growth and PHA formation. Because the residual biomass might also contain some insoluble solids and PHB granules lost in PHB recovery, both cell density and PHB concentration were compared at 24 hours and 48 hours to show the net gains. The benefits of biomass hydrolysates were statistically significant based on the deviations of duplicates. The acid hydrolysates might have two positive effects on microbial PHA formation. First, the hydrolysates promoted cell activity on glucose utilization, giving higher cell densities than the controls in the first 24 hours. This nutritional effect was similar to those of organic nutrients such as yeast extract and peptone, which are widely used in microbial cultures to provide nutrients and growth factors to the cells. A fast cell growth can reduce the cultivation time, resulting in a high PHB productivity. Second, the biomass hydrolysates might also be used as an extra carbon source to generate more cell mass than the controls in 48 hours. This carbon source effect, however, might play a minor role because the cell density did not increase with cell debris load. In fact, too much acid hydrolysates deteriorated the gains as shown in Figure 7. The reason is not clear yet. A load of acid biomass hydrolysates to glucose from 10 to 20 wt%

An acid-base biomass hydrolysates solution containing 48.5 g/L of soluble solids was added into a glucose medium at predetermined percentage of cell debris to glucose from 0 to 40%. The glucose medium without biomass hydrolysates was run in parallel as controls. As shown

**3.4. Utilization of acid hydrolysates in PHB biosynthesis**

seems appropriate for both cell growth and PHB formation.

**3.5. Utilization of acid-base biomass hydrolysates**

**Figure 7.** Effect of acid hydrolysates to glucose loading ratio on cell growth (top) and PHB formation (bottom) in reuse of the residual biomass for PHB production.

in Table 1, the concentrations of both cell mass and PHA content, after 48 hours cultivation, were substantially higher than those of the control. The overall cell growth yield (Yx/s) and PHA formation yield (Yp/s) are calculated from the amounts of cell mass and PHA formed in 48 hours based on the initial concentration of glucose. The relative yields (Yx' and Yp') based on the controls were increased by 100 – 300%. More interestingly, the inhibitory effect of acid biomass hydrolysates was not observed in the use of acid-base hydrolysates, and the load of cell debris to glucose can be increased to about 39 wt%. Most likely, the unknown inhibitors in acid hydrolysates were further decomposed into less inhibitory hydrolysates in the base treatment. Since the acid biomass hydrolysates contained primarily the cytoplasm proteins released from the damaged cells, the soluble proteins might adversely affect cell growth at high concentrations. After being hydrolyzed in base solution into peptides and amino acids, the small molecule hydrolysates become less inhibitory and more usable to the cells.

Generation and Utilization of Microbial Biomass

Hydrolysates in Recovery and Production of Poly(3-hydroxybutyrate) 45

downstream recovery and purification as shown in Figure 1 can generate 31 kg PHB resin and 13 kg acid-base hydrolysis of residual microbial biomass. It is assumed that the acid hydrolysates are not separated, but hydrolyzed sequentially in the base treatment and discharged with the base hydrolysates together. If this amount of residual biomass is reused in next PHB fermentation, the percentage of biomass hydrolysates to glucose is 13% at maximum (13 kg for 100 kg glucose), a moderate load of biomass hydrolysates (Table 1). In real fermentations, more glucose is often added because of the residual glucose in the spent medium. This quick calculation indicates that most of residual biomass discharged from downstream separations can be reused in the next PHA fermentation. In addition to the elimination of a waste stream, the productivity and yields of PHA fermentation can also be

SDS is a popular surfactant used in PHA recovery to disrupt the cells or remove a small amount of hydrophobic impurities from PHB granules [30, 31]. The purity of PHB granules can be increased from 96.4% to above 99% when a small amount of SDS was added in the base treatment. The surfactant left in the base solution, however, may have an adverse effect when the cell debris is reused in microbial PHB fermentation. An acid-base biomass hydrolysates solution containing 48.5 g/L of soluble solids and SDS were added into a glucose medium. The cell debris concentration was kept at 1.94 g/L, and SDS concentration was increased from 0.2 to 0.8 g/L with SDS. Controls without biomass hydrolysates and SDS were run in parallel. Table 2 gives the results of cell growth and PHA formation at different

24 hours 48 hours

Cell mass (g/L)

PHB (wt%)

PHB (wt%)

0 0 2.08 ± 0.02 36.8 ± 1.1 2.7 ± 0.02 44.4 ± 0.6 1.94 0.2 3.69 ± 0.06 49.7 ± 2.1 4.99 ± 0.04 60.6 ± 1.1 1.94 0.4 3.27 ± 0.05 39.6 ± 1.2 4.58 ± 0.04 53.8 ± 0.8 1.94 0.6 2.70 ± 0.04 33.9 ± 1.5 3.51 ± 0.03 50.3 ± 0.7 1.94 0.8 2.59 ± 0.04 17.8 ± 1.3 3.54 ± 0.03 25.8 ± 1.2

**Table 2.** Effect of surfactant SDS and biomass hydrolysates on cell growth and PHB formation

Compared with the controls of no biomass hydrolysates and surfactant, all the cultures containing the acid-base hydrolysates exhibited better cell growth. Particularly, the increase of cell concentration from 24 hours to 48 hours was the new cell mass formed in 24 hours from glucose and cell debris in the presence of SDS. The formation of PHB, however, was deteriorated at high SDS concentrations (0.6-0.8 g/L). At a low or moderate SDS concentrations (0.2-0.4 g/L), the positive effect from biomass hydrolysates was much higher than the negative effect of surfactant. The PHB concentrations, after 48 hours cultivation,

significantly improved.

Biomass hydrolysates (g/L)

**3.7. Effect of SDS in biomass hydrolysates solution**

surfactant levels at 24 and 48 hours, respectively.

Cell mass (g/L)

Note: flask cultures were maintained at 30 oC and 200 rpm in a rotary incubator.

SDS (g/L)

Note: the flask cultures were maintained at 30 oC and 200 rpm in a rotary incubator for 48 hours. The yields of cell mass (Yx/s) and PHB (Yp/s) were based on the initial glucose concentration. The relative yields (Yx' and Yp') are the ratios of yields with hydrolysates to the control without hydrolsyates.

**Table 1.** Effect of acid-base biomass hydrolysates on cell growth and PHB formation

### **3.6. Comparison of three types of cell debris**

In the process of PHB recovery (Figure 1), three types of biomass hydrolysates may be generated, depending on operations: acid, base, and acid-base biomass hydrolysates. They may have different nutrient values or inhibitory effects on cell growth and PHB synthesis. Solutions of three types of biomass hydrolysates were added into a glucose medium for predetermined concentrations of cell debris. Controls without hydrolysates were run in parallel. The ratios of cell densities (g/L) to the controls were compared after 48 hours cultivation as shown in Figure 8. The nutritional value of acid hydrolysates in cell growth is similar to that of base hydrolysates. The nutrient value of acid-base hydrolysates, however, is significantly higher than those of hydrolysates from individual treatment.

**Figure 8.** Comparison of three types of biomass hydrolysates (acid, base, and acid-base) on cell growth in a glucose mineral medium. The relative cell gain is the ratio of cell density to the controls.

Based on an average cell yield (Yx/s = 0.45) of PHB fermentation on glucose [29], 45 kg of cell mass containing 70 wt% of PHB is generated from 100 kg of glucose consumed. A downstream recovery and purification as shown in Figure 1 can generate 31 kg PHB resin and 13 kg acid-base hydrolysis of residual microbial biomass. It is assumed that the acid hydrolysates are not separated, but hydrolyzed sequentially in the base treatment and discharged with the base hydrolysates together. If this amount of residual biomass is reused in next PHB fermentation, the percentage of biomass hydrolysates to glucose is 13% at maximum (13 kg for 100 kg glucose), a moderate load of biomass hydrolysates (Table 1). In real fermentations, more glucose is often added because of the residual glucose in the spent medium. This quick calculation indicates that most of residual biomass discharged from downstream separations can be reused in the next PHA fermentation. In addition to the elimination of a waste stream, the productivity and yields of PHA fermentation can also be significantly improved.

### **3.7. Effect of SDS in biomass hydrolysates solution**

44 Biomass Now – Cultivation and Utilization

Biomass /Glucose (wt%)

ratios of yields with hydrolysates to the control without hydrolsyates.

**3.6. Comparison of three types of cell debris**

0

1

2

**Relative Cell gain**

3

Cell density (g/L)

**Table 1.** Effect of acid-base biomass hydrolysates on cell growth and PHB formation

is significantly higher than those of hydrolysates from individual treatment.

PHB

Yx/s

Yp/s

Yx' Yp'

(g/g)

(g/g)

(wt%)

0.0 0 2.1 ± 0.5 45 ± 1.2 0.18 0.081 1.0 1.0 1.15 9.4 4.0 ± 0.2 55 ± 1.6 0.33 0.18 1.8 2.24 2.30 18.8 4.5 ± 0.3 60 ± 1.7 0.38 0.23 2.1 2.84 4.60 38.8 5.6 ± 0.4 61± 1.5 0.47 0.29 2.6 3.58

Note: the flask cultures were maintained at 30 oC and 200 rpm in a rotary incubator for 48 hours. The yields of cell mass (Yx/s) and PHB (Yp/s) were based on the initial glucose concentration. The relative yields (Yx' and Yp') are the

In the process of PHB recovery (Figure 1), three types of biomass hydrolysates may be generated, depending on operations: acid, base, and acid-base biomass hydrolysates. They may have different nutrient values or inhibitory effects on cell growth and PHB synthesis. Solutions of three types of biomass hydrolysates were added into a glucose medium for predetermined concentrations of cell debris. Controls without hydrolysates were run in parallel. The ratios of cell densities (g/L) to the controls were compared after 48 hours cultivation as shown in Figure 8. The nutritional value of acid hydrolysates in cell growth is similar to that of base hydrolysates. The nutrient value of acid-base hydrolysates, however,

**Figure 8.** Comparison of three types of biomass hydrolysates (acid, base, and acid-base) on cell growth

0123456 **Cell debris (g/L)**

Acid Base Acid-Base

Based on an average cell yield (Yx/s = 0.45) of PHB fermentation on glucose [29], 45 kg of cell mass containing 70 wt% of PHB is generated from 100 kg of glucose consumed. A

in a glucose mineral medium. The relative cell gain is the ratio of cell density to the controls.

Biomass hydrolysates (g/L)

> SDS is a popular surfactant used in PHA recovery to disrupt the cells or remove a small amount of hydrophobic impurities from PHB granules [30, 31]. The purity of PHB granules can be increased from 96.4% to above 99% when a small amount of SDS was added in the base treatment. The surfactant left in the base solution, however, may have an adverse effect when the cell debris is reused in microbial PHB fermentation. An acid-base biomass hydrolysates solution containing 48.5 g/L of soluble solids and SDS were added into a glucose medium. The cell debris concentration was kept at 1.94 g/L, and SDS concentration was increased from 0.2 to 0.8 g/L with SDS. Controls without biomass hydrolysates and SDS were run in parallel. Table 2 gives the results of cell growth and PHA formation at different surfactant levels at 24 and 48 hours, respectively.

Note: flask cultures were maintained at 30 oC and 200 rpm in a rotary incubator.

**Table 2.** Effect of surfactant SDS and biomass hydrolysates on cell growth and PHB formation

Compared with the controls of no biomass hydrolysates and surfactant, all the cultures containing the acid-base hydrolysates exhibited better cell growth. Particularly, the increase of cell concentration from 24 hours to 48 hours was the new cell mass formed in 24 hours from glucose and cell debris in the presence of SDS. The formation of PHB, however, was deteriorated at high SDS concentrations (0.6-0.8 g/L). At a low or moderate SDS concentrations (0.2-0.4 g/L), the positive effect from biomass hydrolysates was much higher than the negative effect of surfactant. The PHB concentrations, after 48 hours cultivation, were 2.4 to 3 g/l, in comparison with 1.2 g/L of the control. The results in Table 2 indicate that the dosage of SDS in PHA recovery should be controlled according to the amount of residual microbial biomass generated. The mass ratio of SDS to biomass hydrolysates should be less than 20% w/w or better at 10% w/w. In a typical PHB recovery process as shown in Figure 1, the amount of SDS used should be less than 2.9 g for 10% of acid-base cell debris or 5.8 g for 20% of acid-base cell debris. The consumption of SDS is therefore 4-8 % of PHB resin produced. It is much lower than the SDS dosages used in the conventional separations [30, 31].

Generation and Utilization of Microbial Biomass

Hydrolysates in Recovery and Production of Poly(3-hydroxybutyrate) 47

[2] Steinbuchel A, Valentin HE (1995) Diversity of bacterial polyhydroxyalkanoic acids.

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### **4. Conclusion**

Residual microbial biomass is an inevitable waste generated in downstream recovery of polyhydroxyalkanoates from microbial cells. With a separation technology based on sequential dissolution of no-PHB cell mass in aqueous solutions, the cell mass separated from the PHB-granules is decomposed and hydrolyzed into small molecule hydrolysates that can be assimilated by microbial cells as nutrients and/or carbon source. A type of biomass hydrolysates generated from continuing treatment in acid and base solutions exhibits the best nutrient value for cell growth and PHA formation. The acid-base hydrolysates contains two major water-soluble components derived from the cell proteins and lipids, respectively. When PHB-producing cells are fed with the hydrolysates in a glucose mineral solution, the cells grow faster and form more biopolyester in comparison with the controls that do not contain the hydrolysates. The glucose-based yields of cell mass and PHA bioplastics are significantly improved. SDS is an efficient surfactant to remove the small amount of hydrophobic residues for high PHB purity, but also a potential inhibitor to microbial PHA formation. When the amount of surfactant is less than 20% of an acid-base biomass hydrolysates, its negative effect is overwhelmed by the nutritional value of hydrolysates. Under these conditions, it is highly possible to reuse most of the residual biomass discharged from PHB recovery in the next microbial PHB fermentation. It therefore eliminates a waste stream from bioplastics production and saves the nutrients with improved PHA productivity and yield.

### **Author details**

Jian Yu, Michael Porter and Matt Jaremko *Hawaii Natural Energy Institute, University of Hawaii at Manoa, Honolulu, Hawaii, USA*

### **Acknowledgement**

The authors acknowledge a support from Bio-On to this work. MP and MJ are graduate students of Molecular Bioscience & Bioengineering at UHM.

### **5. References**

[1] Shuler ML, Kargi F (1992) Bioprocess Engineering: Basic Concepts. New Jersey: Prentice-Hall Inc. pp. 48-50.

[2] Steinbuchel A, Valentin HE (1995) Diversity of bacterial polyhydroxyalkanoic acids. *FEMS Microbiol. Lett.* 128:219-228.

46 Biomass Now – Cultivation and Utilization

improved PHA productivity and yield.

Jian Yu, Michael Porter and Matt Jaremko

Prentice-Hall Inc. pp. 48-50.

students of Molecular Bioscience & Bioengineering at UHM.

**Author details**

**Acknowledgement**

**5. References**

**4. Conclusion**

were 2.4 to 3 g/l, in comparison with 1.2 g/L of the control. The results in Table 2 indicate that the dosage of SDS in PHA recovery should be controlled according to the amount of residual microbial biomass generated. The mass ratio of SDS to biomass hydrolysates should be less than 20% w/w or better at 10% w/w. In a typical PHB recovery process as shown in Figure 1, the amount of SDS used should be less than 2.9 g for 10% of acid-base cell debris or 5.8 g for 20% of acid-base cell debris. The consumption of SDS is therefore 4-8 % of PHB resin produced. It is much lower than the SDS dosages used in the conventional separations [30, 31].

Residual microbial biomass is an inevitable waste generated in downstream recovery of polyhydroxyalkanoates from microbial cells. With a separation technology based on sequential dissolution of no-PHB cell mass in aqueous solutions, the cell mass separated from the PHB-granules is decomposed and hydrolyzed into small molecule hydrolysates that can be assimilated by microbial cells as nutrients and/or carbon source. A type of biomass hydrolysates generated from continuing treatment in acid and base solutions exhibits the best nutrient value for cell growth and PHA formation. The acid-base hydrolysates contains two major water-soluble components derived from the cell proteins and lipids, respectively. When PHB-producing cells are fed with the hydrolysates in a glucose mineral solution, the cells grow faster and form more biopolyester in comparison with the controls that do not contain the hydrolysates. The glucose-based yields of cell mass and PHA bioplastics are significantly improved. SDS is an efficient surfactant to remove the small amount of hydrophobic residues for high PHB purity, but also a potential inhibitor to microbial PHA formation. When the amount of surfactant is less than 20% of an acid-base biomass hydrolysates, its negative effect is overwhelmed by the nutritional value of hydrolysates. Under these conditions, it is highly possible to reuse most of the residual biomass discharged from PHB recovery in the next microbial PHB fermentation. It therefore eliminates a waste stream from bioplastics production and saves the nutrients with

*Hawaii Natural Energy Institute, University of Hawaii at Manoa, Honolulu, Hawaii, USA*

The authors acknowledge a support from Bio-On to this work. MP and MJ are graduate

[1] Shuler ML, Kargi F (1992) Bioprocess Engineering: Basic Concepts. New Jersey:

[18] Fiorese ML, Freitas F, Pais J, Ramos AM, de Aragao GMF, Reis MAM (2009) Recovery of polyhydroxybutyrate (PHB) from *Cuprivavidus necator* biomass by solvent extraction with 1,2-propylene carbonate. *Eng. Life Sci*. 9:454-461.

**Chapter 3**

© 2013 Bruckman et al., licensee InTech. This is an open access chapter distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

© 2013 Bruckman et al., licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

**Considerations for Sustainable**

*Quercus***-Dominated Forest Ecosystems**

Viktor J. Bruckman, Shuai Yan, Eduard Hochbichler and Gerhard Glatzel

Our current energy system is mainly based on carbon (C) intensive metabolisms, resulting in great effects on the earth's biosphere. The majority of the energy sources are fossil (crude oil, coal, natural gas) and release CO2 in the combustion (oxidation) process which takes place during utilization of the energy. C released to the atmosphere was once sequestered by biomass over a time span of millions of years and is now being released back into the atmosphere within a period of just decades. Fossil energy is relatively cheap and has been fuelling the world economy since the industrial revolution. To date, fossil fuel emissions are still increasing despite a slight decrease in 2009 as a consequence of the world's economic crisis. Recently, the increase is driven by emerging economies, from the production and international trade of goods and services [1]."If we don´t change direction soon, we´ll end up where we´re heading" is the headline of the first paragraph in the executive summary of the World Energy Outlook 2011 [2]. It unfortunately represents systematic failure in combating climate change and the emphatic introduction of a "green society", leaving the fossil age behind. Certainly such far reaching transformations would take time, but recovery of the world economy since 2009, although uneven, again resulted in rising global primary energy demands [2]. It seems that more or less ambitious goals for climate change prevention are only resolved in phases of a relatively stable economy. Atmospheric carbon dioxide (CO2) is the second most important greenhouse warming agent after water vapour, corresponding to 26% and 60% of radiative forcing, respectively [3]. Together with other greenhouse gases (GHG's) (e.g. methane (CH4), nitrous oxide (N2O) or ozone (O3)) they contribute to anthropogenic global warming. The industrialization has been driven by fossil sources of energy, emerging in the 17th and 18th century in England as a historical

**Biomass Production in**

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/53518

singularity, but soon spreading globally.

**1. Introduction**

## **Considerations for Sustainable Biomass Production in**  *Quercus***-Dominated Forest Ecosystems**

Viktor J. Bruckman, Shuai Yan, Eduard Hochbichler and Gerhard Glatzel

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/53518

### **1. Introduction**

48 Biomass Now – Cultivation and Utilization

[18] Fiorese ML, Freitas F, Pais J, Ramos AM, de Aragao GMF, Reis MAM (2009) Recovery of polyhydroxybutyrate (PHB) from *Cuprivavidus necator* biomass by solvent extraction

[19] Kapritchkoff FM, Viotti AP, Alli RCP, Succolo M, Pradella JGC, Maiorano AE, Miranda EA, Bonomi A (2006) Enzymatic recovery and purification of polyhydroxybutyrate

[20] Yu J, Chen L (2006). Cost effective recovery and purification of polyhydroxyalkanoates

[21] Yu J, Plackett D, Chen LXL (2005). Kinetics and mechanism of the monomeric products from abiotic hydrolysis of poly(R)-3-hydroxybutyrate under acidic and alkaline

[22] Yu, J (2009). Recovery and purification of polyhydroxyalkanoates from PHA-containing

[23] Bradford M (1976) A rapid and sensitive method for the quantization of microgram quantities of protein utilizing the principle of protein-dye binding. *Anal. Biochem*.

[24] Hesselmann RPX, Fleischmann T, Hany R, Zehnder AJB (1999) Determination of polyhydroxyalkanoates in activated sludge by ion chromatographic and enzymatic

[25] Chittur KK (1998). FTIR/ATR for protein adsorption to biomaterial surfaces. *Biomaterials*

[26] Kansiz M, Billman-Jacobe H, McNaughton D (2000) Quantitative determination of the biodegradable polymer poly(-hydroxybutyrate) in a recombinant *Escherichia coli* strain by use of mid-infrared spectroscopy and multivariative statistics. *Appl. Environ.*

[27] Bloembergen S, Holden DA, Hamer GK, Bluhm TL, Marchessault RM (1986). Studies of composition and crystallinity of bacterial poly(-hydroxybutyrate-co--

[28] Barham PJ, Keller A, Otun EL, Holmes PA (1984) Crystallization and morphology of a

[29] Kim BS, Lee SC, Lee SY, Chang HN, Chang YK, Woo SI (1994). Production of poly(3 hydroxybutyraic acid) by fed-batch culture of *Alcaligenes eutrophus* with glucose

[30] Chen Y, Xu Q, Yang H, Gu G (2001). Effects of cell fermentation time and biomass drying strategies on the recovery of poly-3-hydroxyalkanoates from *Alcaligenes*

*eutrophus* using a surfactant-chelate aqueous system. *Process Biochem*. 36:773-779. [31] Strazzullo G, Gambacorta A, Vella FM, Immirzi B, Romano I, Calandrelli V, Nicolaus B, Lama L (2008). Chemical-physical characterization of polyhydroxyalkanoates recovery by means of a simplified method from cultures of *Halomonas campaniensis*. *World J.*

bacterial thermoplastic – Poly-3-hydroxybutyrate. *J. Mater Sci*. 19:2781-2794.

with 1,2-propylene carbonate. *Eng. Life Sci*. 9:454-461.

produced by *Ralstonia eutropha*. *J. Biotechnol*. 122:453-462.

conditions. *Polym. Degrad. Stabil.* 89:289-299.

methods. *J. Microbiol. Methods* 35:111-119.

hydroxyvalerate). *Macromolecules* 19:2865-2871.

concentration control. *Biotech. Bioeng*. 892-898.

cell mass. *US Patent 7,514,525*.

72:248-254.

19:357-369.

*Microbiol*. 66:3415-3420.

*Microb. Biot.* 24:1513-1519.

by selective dissolution of cell mass. *Biotechnol. Progr.* 22:547-553.

Our current energy system is mainly based on carbon (C) intensive metabolisms, resulting in great effects on the earth's biosphere. The majority of the energy sources are fossil (crude oil, coal, natural gas) and release CO2 in the combustion (oxidation) process which takes place during utilization of the energy. C released to the atmosphere was once sequestered by biomass over a time span of millions of years and is now being released back into the atmosphere within a period of just decades. Fossil energy is relatively cheap and has been fuelling the world economy since the industrial revolution. To date, fossil fuel emissions are still increasing despite a slight decrease in 2009 as a consequence of the world's economic crisis. Recently, the increase is driven by emerging economies, from the production and international trade of goods and services [1]."If we don´t change direction soon, we´ll end up where we´re heading" is the headline of the first paragraph in the executive summary of the World Energy Outlook 2011 [2]. It unfortunately represents systematic failure in combating climate change and the emphatic introduction of a "green society", leaving the fossil age behind. Certainly such far reaching transformations would take time, but recovery of the world economy since 2009, although uneven, again resulted in rising global primary energy demands [2]. It seems that more or less ambitious goals for climate change prevention are only resolved in phases of a relatively stable economy. Atmospheric carbon dioxide (CO2) is the second most important greenhouse warming agent after water vapour, corresponding to 26% and 60% of radiative forcing, respectively [3]. Together with other greenhouse gases (GHG's) (e.g. methane (CH4), nitrous oxide (N2O) or ozone (O3)) they contribute to anthropogenic global warming. The industrialization has been driven by fossil sources of energy, emerging in the 17th and 18th century in England as a historical singularity, but soon spreading globally.

© 2013 Bruckman et al., licensee InTech. This is an open access chapter distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 Bruckman et al., licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Today, our economies still rely on relatively cheap sources of fossil energy, mainly crude oil and natural gas, and consequently emitting as much as 10 PG C per year in 2010 [4]. The Mauna Loa Observatory in Hawaii carries out the most comprehensive and longest continuous monitoring of atmospheric CO2 concentration. It publishes the well-known Keeling Curve, representing the dynamic change since 1958. Observing the Keeling Curve, one can easily recognize the seasonal variability which is directly triggered by CO2 uptake of vegetation (biomass) in the northern hemisphere during the vegetation period and secondly, which is even more important in terms of global change, a steady increase of CO2 concentration from 315 ppmv in 1958 to 394 ppmv in March 2012 [5]. Earlier concentrations could still be derived from air occluded in ice cores. Neftel et al. [6] presents accurate gas concentration measurements for the past two centuries. However, the theoretical knowledge of the warming potential of CO2 in the atmosphere evolved in the late 19th century when a theory of climate change was proposed by Plass [7], pointing out the "influence of man's activities on climate" as well as the CO2 exchange between oceans and atmosphere and subsequent acidification. He highlighted the radiative flux controlled by CO2 in the 12 to 18 micron frequency interval, agreeing with a number of studies published in the forthcoming decades, e.g. Kiehl and Trenberth [3]. In order to understand the fate of anthropogenic CO2 emissions, research soon focussed on estimating sources and sinks as well as their stability, since it was obvious that the atmospheric concentrations did not rise at the same magnitude as emissions. Available numbers on current fluxes are principally based on the work of Canadell et al., [8] and Le Quéré et al., [1]. In their studies, it is emphasized that the efficiency of the sinks of anthropogenic C is expected to decrease. Sink regions (of ocean and land) could have weakened, source regions could have intensified or sink regions could have transitioned to sources [8]. Another explanation might be the fact that the atmospheric CO2 concentration is increasing at a higher rate than the sequestration rate of sinks [1]. Moreover, CO2 fertilization on land is limited as the positive effect levels off and the carbonate concentration which buffers CO2 in the ocean steadily decreases according to Denman, K.L. et al. [1]. Fossil fuel combustion and land use change (LUC) are the major sources for anthropogenic C emissions (Figure 1). Land use change is usually associated with agricultural practices and intensified agriculture triggers deforestation in developing countries [9] and consequently causes additional emissions.

Considerations for Sustainable Biomass Production in *Quercus*-Dominated Forest Ecosystems 51

2.6±1.0 **26%**

2.4±0.5 **24%**

5.0±0.2 **50%**

**Figure 1.** The fate of anthropogenic CO2 emissions in 2010, showing sources (left) and sinks (right). Presented numbers are Pg C yr-1. The values for 2010 were presented at the Planet under Pressure 2012

combustion\* Atmosphere Ocean Terrestrial

Biomass could play a significant role in the renewable energy mix. It is a feedstock for bioenergy production and currently thermal utilization (combustion) is by far the most important conversion process, but research activities are focussed on a range of different processes. This includes, for instance, the Fischer-Tropsch synthesis where any kind of biomass may be used as feedstock to produce liquid biofuels. This process is known as biomass – to – liquid (BtL). Research is pushed by national and international regulations (e.g. the EU's 2020 bioenergy target) and commitments, as a climate change mitigation strategy.

This chapter focuses on aspects of sustainable woody biomass production in *Quercus* dominated forest ecosystems with emphasis on different silvicultural management systems. Short rotation woody crops (SRWC), coppice with standards (CS), high forest (HF) and Satoyama are characterized according to their biomass potential and sustainability considerations. CS and HF are directly compared based on our own research and links to similar systems (SRWC and Satoyama) are drawn in order to provide a holistic view of the current topic. The chapter aims at providing an interdisciplinary view on biomass production in forest ecosystems, considering impacts on C and nutrient metabolism as well as other effects (e.g. biodiversity, technical, silvicultural and cultural issues). Considerations for sustainable biomass production in *Quercus* dominated forest ecosystems are presented

Terrestrial C sequestration accounts for approximately one quarter of the three main sinks as indicated in Figure 2, where forests contribute the largest share. An intact terrestrial sink might be more important in the future in terms of mitigating climate change, since the ocean

conference in London [4].

\*includes cement production and flaring.

0.9±0.7 **9%**

Land use change Fossil fuel

9.1±0.5 **91%**

for each management system.

**1.1. Biomass production and carbon sequestration**

Another consideration is the availability of fossil fuel, which is limited by the fact that it is a non-renewable and therefore finite resource. Since the fossil energy system is based on globally traded sources with centralized structures, the vulnerability to disturbance is high. Recent examples of price fluctuations caused by political crises or other conflicts in producing countries or along transportation lines demonstrate potential risks. Moreover, a shift towards alternative energy sources and a decentralization of the energy system may contribute to system resilience and create domestic jobs. It prevents capital outflow to unstable political regimes and it helps to protect the environment not only by reducing GHG emissions, but also by reducing impacts of questionable methods of extracting fossil sources of energy (e.g. tar sands exploitation, fracking etc.).

**Figure 1.** The fate of anthropogenic CO2 emissions in 2010, showing sources (left) and sinks (right). Presented numbers are Pg C yr-1. The values for 2010 were presented at the Planet under Pressure 2012 conference in London [4].

\*includes cement production and flaring.

50 Biomass Now – Cultivation and Utilization

countries [9] and consequently causes additional emissions.

sources of energy (e.g. tar sands exploitation, fracking etc.).

Another consideration is the availability of fossil fuel, which is limited by the fact that it is a non-renewable and therefore finite resource. Since the fossil energy system is based on globally traded sources with centralized structures, the vulnerability to disturbance is high. Recent examples of price fluctuations caused by political crises or other conflicts in producing countries or along transportation lines demonstrate potential risks. Moreover, a shift towards alternative energy sources and a decentralization of the energy system may contribute to system resilience and create domestic jobs. It prevents capital outflow to unstable political regimes and it helps to protect the environment not only by reducing GHG emissions, but also by reducing impacts of questionable methods of extracting fossil

Today, our economies still rely on relatively cheap sources of fossil energy, mainly crude oil and natural gas, and consequently emitting as much as 10 PG C per year in 2010 [4]. The Mauna Loa Observatory in Hawaii carries out the most comprehensive and longest continuous monitoring of atmospheric CO2 concentration. It publishes the well-known Keeling Curve, representing the dynamic change since 1958. Observing the Keeling Curve, one can easily recognize the seasonal variability which is directly triggered by CO2 uptake of vegetation (biomass) in the northern hemisphere during the vegetation period and secondly, which is even more important in terms of global change, a steady increase of CO2 concentration from 315 ppmv in 1958 to 394 ppmv in March 2012 [5]. Earlier concentrations could still be derived from air occluded in ice cores. Neftel et al. [6] presents accurate gas concentration measurements for the past two centuries. However, the theoretical knowledge of the warming potential of CO2 in the atmosphere evolved in the late 19th century when a theory of climate change was proposed by Plass [7], pointing out the "influence of man's activities on climate" as well as the CO2 exchange between oceans and atmosphere and subsequent acidification. He highlighted the radiative flux controlled by CO2 in the 12 to 18 micron frequency interval, agreeing with a number of studies published in the forthcoming decades, e.g. Kiehl and Trenberth [3]. In order to understand the fate of anthropogenic CO2 emissions, research soon focussed on estimating sources and sinks as well as their stability, since it was obvious that the atmospheric concentrations did not rise at the same magnitude as emissions. Available numbers on current fluxes are principally based on the work of Canadell et al., [8] and Le Quéré et al., [1]. In their studies, it is emphasized that the efficiency of the sinks of anthropogenic C is expected to decrease. Sink regions (of ocean and land) could have weakened, source regions could have intensified or sink regions could have transitioned to sources [8]. Another explanation might be the fact that the atmospheric CO2 concentration is increasing at a higher rate than the sequestration rate of sinks [1]. Moreover, CO2 fertilization on land is limited as the positive effect levels off and the carbonate concentration which buffers CO2 in the ocean steadily decreases according to Denman, K.L. et al. [1]. Fossil fuel combustion and land use change (LUC) are the major sources for anthropogenic C emissions (Figure 1). Land use change is usually associated with agricultural practices and intensified agriculture triggers deforestation in developing

Biomass could play a significant role in the renewable energy mix. It is a feedstock for bioenergy production and currently thermal utilization (combustion) is by far the most important conversion process, but research activities are focussed on a range of different processes. This includes, for instance, the Fischer-Tropsch synthesis where any kind of biomass may be used as feedstock to produce liquid biofuels. This process is known as biomass – to – liquid (BtL). Research is pushed by national and international regulations (e.g. the EU's 2020 bioenergy target) and commitments, as a climate change mitigation strategy.

This chapter focuses on aspects of sustainable woody biomass production in *Quercus* dominated forest ecosystems with emphasis on different silvicultural management systems. Short rotation woody crops (SRWC), coppice with standards (CS), high forest (HF) and Satoyama are characterized according to their biomass potential and sustainability considerations. CS and HF are directly compared based on our own research and links to similar systems (SRWC and Satoyama) are drawn in order to provide a holistic view of the current topic. The chapter aims at providing an interdisciplinary view on biomass production in forest ecosystems, considering impacts on C and nutrient metabolism as well as other effects (e.g. biodiversity, technical, silvicultural and cultural issues). Considerations for sustainable biomass production in *Quercus* dominated forest ecosystems are presented for each management system.

### **1.1. Biomass production and carbon sequestration**

Terrestrial C sequestration accounts for approximately one quarter of the three main sinks as indicated in Figure 2, where forests contribute the largest share. An intact terrestrial sink might be more important in the future in terms of mitigating climate change, since the ocean

sink is expected to decrease. Our current forests are capable of sequestering ~2.4 Pg C yr-1 (of 2.6 Pg in total), when excluding tropical land-use change areas [10]. Sequestration of C in forests is controlled by environmental conditions, disturbance and management. However, forests can principally act as a source or sink of C, depending on the balance between photosynthesis and respiration, decomposition, forest fire and harvesting operations. On both European and global scales, forests were estimated to act as sinks on average over the last few decades [11, 12].

Considerations for Sustainable Biomass Production in *Quercus*-Dominated Forest Ecosystems 53

subsoil, but considerable amounts can still be found if one not only analyses the topsoil layer as recommended by a number of authors, e.g. by Diochon et al*.* [18]. In contrast, the radiocarbon age of the topsoil may range from less than a few decades [17] to months if considering freshly decomposed organic matter. The reasons for the relatively high age in subsoil horizons are not clear, but unfavourable conditions for soil microbial diversity and strong association of C with mineral surfaces (organo-mineral interactions with clay minerals) might be an explanation [16]. There are, in principal, two pathways for sequestering C in forest soils. Forest litter consists of leaves, needles and woody debris; such as branches, bark and fruit shells which accumulate on the surface (L and F layer). Soil macrofauna degrades it until it becomes part of the organic matter (OM) where it is impossible to recognize its original source (H layer). Parts of it are translocated into deeper horizons by bioturbation (e.g. earthworms) or remain on the surface, to be further degraded by soil microorganisms while organic matter becomes mobile in the form of humic acids and subsequently being mineralized at a range of negatively charged surfaces (humus, clay minerals). The second pathway is through root turnover and rhizodeposition (= excretion of root exudates). Matthews and Grogan [19] and subsequently Grogan and Matthews [20] parameterized their models with values of between 50 and 85% of C from the fine root pool which is lost to the SOC pool on an annual basis, depending on species composition and management. This assumption is consistent with another study where 50% of the living fine roots were assumed to reflect real values [21]. There is evidence that C derived from root biomass [22] and mycorrhizal hyphal turnover [23] might be the most important source for SOC pools rather than from litter decomposition. Since fine root turnover is species specific [24], it could be controlled to some extent by species composition at management unit levels. A further question for a forest owner in terms of carbon sequestration is which management option to choose while still being able to produce products and generate income. Despite the fact that unmanaged forests hold the highest C pools, it was commonly believed that aggrading forests reach a maximum sequestration and it is reduced in old growth forests, where photosynthesis and autotrophic respiration are close to offsetting each other. This is contrasts a number of studies, pointing out that even unmanaged forests at a late successional development stage could still act as significant carbon sinks [25]. The authors of another study claim that advanced forests should not be neglected from the carbon sequestration discussion a priori [26] and they should be left intact since they will lose much of their C when disturbed [27]. Therefore, managing forests implies a trade-off between maximum C sequestration and provision of goods, such as timber and biomass for energetic utilization. While the highest amounts of carbon could be sequestered in unmanaged forests [27, 28], C sequestration through forest management can be a cost-effective way to reduce atmospheric CO2, despite limited quantities, due to biological limitations and societal constraints [29]. This is in general agreement with the conclusion of Wiseman's [30] dissertation, who argues that there is potential for additional C uptake depending on forest management, but the effect is short-term until a new equilibrium in C stocks is reached and

also argues that the effect may be limited.

The most important process of net primary production is achieved by photosynthesis, which is the chemical transformation of atmospheric CO2 and water from the soil matrix into more complex carbohydrates and long chain molecules to build up cellulose, which is found in cell walls of woody tissue as well as hemicellulose and lignin. The C remains in the woody compound either until it is degraded by microorganisms, which use the C as source of energy, or until oxidation takes place (e.g. burning biomass, forest fire). In both cases, it becomes part of atmospheric CO2 again. A certain share, controlled mainly by climatic conditions [13], enters the soil pool as soil organic carbon (SOC). The ratio between aboveground and belowground pools depends on the current stand age, forest management and climate. In temperate managed forests, SOC stocks are typically similar to the aboveground stocks [14], which is confirmed in our own research [13]. SOC (and in particular O-horizon) stocks are typically higher in boreal forests and in high elevation coniferous forests as a consequence of reduced microbial activity and much lower in tropical environments. O-layer C pools are especially sensitive to changes in local climate. A traditional forest management regime in Austrian montane spruce forests is clear-cutting, typically from the top of a hill to the valley to facilitate cable skidding. An abrupt increase of radiative energy and water on the soil surface creates favourable conditions for soil microorganisms and a great amount of C stored in the O-layer will be released to the atmosphere by heterotrophic respiration. Other GHG's, such as N2O are eventually emitted under moist and reductive conditions as excess nitrogen is removed by lateral water flows. Unfortunately, this effect is likely to happen on a large-scale where massive amounts of C might be released as the global temperature rises and thawing permafrost induces C emissions [15], potentially creating a strong feedback cycle, further accelerating global warming. However, Don et al. [14] found that SOC pools were surprisingly stable after a major disturbance (wind throw event), indicating low short-term vulnerability of forest floor and upper mineral horizons. They explained their findings with herbaceous vegetation and harvest residues, taking over the role of litter C input. The study covers a time-span of 3.5 years which might be too short to observe soil C changes. Likewise, we did not observe significant C stock decrease in our youngest sample plot of the coppice with standards (CS) chronosequence [13]. We expect that the N dynamics might have a profound influence on C retention and the impact of disturbance on SOC pools depends on environmental conditions. Successful long-term sequestration in terms of climate change mitigation is therefore only achieved if C becomes part of the recalcitrant fraction in the subsoil, which is typically between 1 000 and 10 000 years old [16, 17]. The C concentration is lower in the

last few decades [11, 12].

sink is expected to decrease. Our current forests are capable of sequestering ~2.4 Pg C yr-1 (of 2.6 Pg in total), when excluding tropical land-use change areas [10]. Sequestration of C in forests is controlled by environmental conditions, disturbance and management. However, forests can principally act as a source or sink of C, depending on the balance between photosynthesis and respiration, decomposition, forest fire and harvesting operations. On both European and global scales, forests were estimated to act as sinks on average over the