Research Article
Influence of Autoclaving of Shrimp Shells on Proteinase Enzyme Production and Protein Hydrolysis by Aspergillus niger
Nesreen Mahmoud and Abdelkader Ghaly*
Corresponding Author: Professor A. E. Ghaly, Department of Process Engineering and Applied Science, Faculty of Engineering, Dalhousie University, Halifax, Nova Scotia, Canada; Tel::902-494-6014; Email: abdel.ghaly@dal.ca
Received: March 26, 2015; Accepted: April 6, 2015; Published: April 14, 2015;
Citation: Mahmoud N & Ghaly A (2015) Influence of Autoclaving of Shrimp Shells on Proteinase Enzyme Production and Protein Hydrolysis by Aspergillus niger. International Journal of Bioprocess and Biotechnological Advancements, 1(1): 1-17
Copyrights: ©2015 Mahmoud N, Ghaly A. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are cr
 


The aim of the study was to evaluate the ability of the fungus Aspergillus niger to carry out the deproteinization process of a carbon supplemented shrimp shells and to study the effect of autoclaving the shrimp shells on the performance characteristics of the deproteinization process. The temperatures of the shrimp shells and the exhaust gas declined during the lag period as the heat losses from the bioreactor were higher than the heat generated by metabolic activities. Once the exponential growth phase started, the temperatures of the shrimp shells and exhaust gas and CO2 concentration increased reaching maximum values of 37.5 and 33.6ºC, 28.9 and 27.9ºC and 0.37% and 0.32% for the runs with autoclave and intact shrimp shells, respectively. A strong correlation between the concentration of carbon dioxide in the exhaust gas and the temperature of the shrimp shells was obtained.  The pH of the shrimp shells initially decreased with time reaching 7.05-7.34 and then increased reaching 7.35-8.16. The decrease in the pH was due to acid protease production while the increase in the pH was due to the buffering capacity of the calcium carbonate released from the shrimp shells and the production of ammonium nitrogen. The galactose concentration and moisture content decreased gradually over time reaching about 1.48% and 2.67% and 25.54% and 26.72% at the end of the experiment for the autoclaved and unautoclaved shells, respectively. The protease activity of the autoclave shells was significantly higher (1.7 fold) than that of the unautoclaved shells. The deproteinization efficiencies of autoclaved and unautoclaved shells were 33.23% and 25.62% for, respectively. The reduction in the protein concentration at the end of the deproteinization process did not correspond to the increase in the protease activity. The low deproteinization efficiency observed in this study could be due to the high initial pH of the shrimp shells and the high temperature and the low moisture content observed during the deproteinization process that might have interfered with protease synthesis and/or activity. The low deproteinization attained with unautoclaved shells might be due to the growth of the indigenous microorganisms in the shrimp shells which might affect the growth and metabolic activity of A. niger. The chitin concentration increased from 16.56 to 22.68% and 20.93% over the course of the deproteinization process for the autoclaved and unautoclaved shrimp shells, respectively. The spent shrimp shells had a pale pink-orange color with some tan patches. The pink-orange color was an indication of the presence of pigments.  The extent of shrimp shell agglomeration and presence of white precipitant were much higher in case of unautoclaved shrimp shells. Unidentified microorganisms were found in both the autoclaved and unautoclaved shells with larger population in the case of unautoclaved shells.

 

Keywords
Shrimp shells, Protein, Chitin, Minerals, Deproteinization, Aspergillus. niger, Protease activity, Galactose, Temperature, pH, Moisture content, Autoclaving.

INTRODUCTION

The shell fish processing industry generates large amounts of solid wastes which have become a major environmental concern to producing countries worldwide due to the high perishability and the bulky nature of the waste material [1, 2]. Northern Pink Shrimp (Pandalus borealis) is commonly fished in the North Atlantic both on the East Coast of Canada and the West Coast of Norway. These shrimp are caught as part of the offshore Northern Shrimp Fishery by a vessel owned by Clearwater Fine Foods Inc. and then individually quick-frozen on board the vessel.  Upon arrival at the cooking/peeling plant, they are cooked in boiling salt water for 10 minutes and then sent to automated peeling machines where the shell and meat portions were separated. This species has a mean length of 22-25 mm at maturity [3] and the processing discards of these shrimp account for up to 80% of its original weight [4]. The total landing of Northern shrimp in Eastern Canada in 2013 was 185974 tonnes [5] which resulted in 148000 tonnes of waste, most of which is dumped in the ocean or landfills.


However, shellfish waste is a rich source of chitin, protein, pigments and flavor compounds but chitin is the most economically valuable component of the waste material [2]. Chitin and its derivatives have been used in many industrial applications including water treatment, pulp and paper, biomedical and therapies devices, cosmetics, biotechnology, agriculture, food science and membrane technology [6,7].


The traditional method of chitin extraction from crustacean waste involves the use of strong acid (HCl) for demineralization and strong alkali (NaOH) for deproteinization [1,2,8,9]. This method yields chitin with variable and inconsistent physiochemical properties, produces wastewater that creates environmental problems, wastes other valuable components in the waste and is costly. Therefore, a less expensive, more efficient and environmentally friendly biological extraction method for chitin from crustacean waste is needed. Several researchers used organic acids for demineralization [1,10-12] and proteolytic microorganisms [8,13-19] or purified proteolytic enzymes [18] for deproteinization of shrimp shells. However, deproteinization by living microbes is more effective and less costly because they provide a gradual increase of protease throughout the process [8,19]. Therefore, chitin extraction using microorganisms is a good alternative to the harsh chemical treatment, less expensive and preserves the natural state of the biopolymer. Aspergillus niger is an important industrial microorganism that is capable of producing proteolytic enzymes [20-23]. It also contains up to 42.0% chitin of the dry weight of the fungal cell wall; the cell wall weight is 20-40% of the dry cell weight [24]. Thus, A. niger has a great potential for deproteinization of crustacean waste while the chitin in the cell wall of the fungi can be considered an additional source of chitin[19]. 


OBJECTIVES

The aim of the research was evaluate the ability of the fungus species Aspergillus niger to produce protease enzyme and carry out the deproteinization process of ground shrimp shells supplemented with galactose as a carbon source and to study the effect of autoclaving the shrimp shells on the performance characteristics of the deproteinization process.


MATERIALS AND METHODS

Experimental Apparatus

Especially designed experimental equipment with a rotary drum bioreactors were used in this study. The experimental setup (Figure 1) consisted of a main frame, three drum bioreactors, an aeration system and a data acquisition system.


The main frame was made of two polyvinyl chloride (PVC) rectangular sheets (13 mm thick) and two hexagon stainless steel sheets (3 mm thick). One of the PVC sheets (560 × 460 mm) was used as a base and the other one (560 × 380 mm) was fixed vertically on the base. The two hexagon stainless steel sheets were fixed to the two PVC sheets by means of stainless steel screws. The main frame held the drum bioreactors, the pressure regulator, the flow meters, the inlet air and exhaust gas manifold, the thermocouple wires, the mixing motor along with the transmission system and the switch, tubing and sampling ports. 


Three 1.8 L drum bioreactors with mixing motors and transmission system were used. Each drum bioreactor (Figure 2) consisted of a removable inner stainless steel mesh (aperture of 1.5 mm) which was used as a lining for an outer stainless steel horizontally rotating basket of 88 mm diameter and 292 mm length. One stainless steel plate, with a drilled hole for sampling, was used  to  close  one  end  of  the  rotating  basket.  The other end was left opened for charging and cleaning purposes and was designed so that it can be recessed and secured into a rotating disc after charging the reactor. An outer casing made from a Plexiglas cylinder of 12.5 mm diameter was installed for each bioreactor. The Plexiglas cylinder was recessed and secured into the main frame from one end by six stainless steel screws. The other end of the Plexiglas cylinder was covered by a removable circular Plexiglas plate and was recessed and secured by six stainless steel screws and wing nuts. A rubber gasket lining (O-ring, 2.5 mm thick) was used at both ends of the Plexiglas cylinder to provide an air tight seal. A hole was drilled through the cylinder wall for the release of the exhaust gas. The rotating discs were connected to a motor (Synchronous Motor, 20-34245G-24007, Xerox127P1292/B, Sigma Instruments Inc., Braintree, Massachusetts,  USA) through a transmission system.


Air was supplied continuously at the required flow rate inside each drum bioreactor from the laboratory air supply. The air passed first through a pressure regulator (ARO, Model no. 129125-510, Bryan, Ohio, USA) in order to regulate the air pressure around 5 kPa and then through a 1 L humidifier which contained 0.75 L sterilized distilled water. The humidified air was passed through a bacterial filter and then through a flow meter (No. 60648, Cole-Parmer Instrument Co., Illinois, USA) and finally introduced into the bioreactor through a small perforated stainless steel tube that ran along the center of the basket. The aeration tube was fixed through the center of the rotating disc and remains stationary while the basket is rotating. The air inlet sampling port was placed right after the bacterial filter whereas the three exhaust gas sampling ports were located on the exhaust tubes. Each sampling port was made of a rubber septum. The three exhaust gas tubes were connected to a manifold and the exhaust gas was bubbled through a small container of water in order to create a slight gas pressure in the bioreactors. 


Eleven T-type thermocouples (Thermo Electric Ltd., Brampton, Ontario, Canada) were used to measure the temperature during the course of the fermentation process. Two thermocouples were threaded through the aeration tube of each bioreactor and used to measure the temperature of the material inside the bioreactor. The other five thermocouples were used to measure the ambient temperature (1), inlet air temperature (1) and exhaust gas temperature of each bioreactor (3). The temperature data were monitored and stored using a data acquisition system  which  consisted  of  a  master  unit  (Multiscan  1200,  Omega,  and  Stamford,  CT),  a thermocouple/volt scanning card (MTC/24, Omega, Stamford, Connecticut, USA), software (Tempview, Omega, Stamford, Connecticut, USA) and a personal computer. 


Microorganisms

The fungus Aspergillus niger (Figure 3) was chosen for this study for two reasons: (a) its ability to produce acid protease and (b) the presence of chitin in its cell wall. The genus Aspergillus is characterized by a well-developed foot cell at the base of the conidiophore. The colony consists of colorless mycelium from which conidiophore arise. The spores develop the black color and the powdery appearance of the colony surface [25].

Aspergillus niger (ATCC 16513) was obtained from the American Type Culture Collection (Rockville, Maryland, USA). The freeze dried culture was revived in 6 mL of 0.1% sterilized peptone solution, which was prepared by dissolving 1 g bacto-peptone (Difco, Detroit, Michigan, USA) in 1 L deionized-distilled water and then sterilized in an autoclave (Model No. STM-E, Market Forge Sterilmatic, New York, USA) at 121 ˚C and 103.4 kPa for 30 minutes. The rehydrated culture was kept in the peptone solution in a capped test tube for 24 hours at room temperature (22 ºC). One mL of the rehydrated A. niger was transferred to each of three test tubes containing 9 mLpotato dextrose broth (PDB), which contained infusion from 200 g potatoes (4 g/L) and 20 g/L Bacto dextrose. The test tubes were kept tightly capped for 48 h at room temperature (22 ºC) and then stored in the fridge at 4 °C and subcultured when needed.

A spore stock suspension was obtained by growing the fungus on Czapek’s agar, (which contained 30.00 g/L saccharose, 2.00 g/L sodium nitrate, 1.00 g/L dipotassium phosphate, 0.50 g/L magnesium sulfate, 0.50 g/L potassium chloride, 0.01 g/L ferrous sulfate and 15.00 g/L agar) at room temperature (22 ºC) for 4 days. The conidia were harvested from the surface by adding sterilized deionized distilled water containing 0.01% (v/v) Tween 80 and gently scraping the surface with a sterile spatula. The Tween 80 was prepared by dissolving 0.1 mL Tween 80 in 1 L distilled deionized water and then autoclaving (Model No. STM-E, Market Forge Sterilmatic, New York, USA) at 121˚ C and 103.4 kPa for 30 minutes. The spore concentration was determined using direct standard plate count method according to the procedures described in the Standard Method for the Examination of Dairy Products [26]. The prepared suspension was stored in the refrigerator at about 4 ºC until needed.

 

Shrimp Shells

The shells of the Northen Pink Shrimp (Pandalus borealis) were obtained from a shell processing plant in Mulgrave, owned by Ocean Nutrition Ltd. of Bedford, Nova Scotia. The shrimp shells were stored at about -25 ºC in the Biotechnology Laboratory till needed. The shrimp shells were dried and then ground using a conventional food processor (General Electric Company, Wal-Mart Canada Corporation, Mississauga, Ontario, Canada). The ground shrimp shells were autoclaved (Model No. STM-E, Market Forge Sterilmatic, New York, USA) at 121˚C and 103.4 kPa for 45 min before use. Table 1 shows some of the characteristics of the shrimp. Table 2 shows the particle size distribution of the ground and intact shrimp shells.


Experimental Protocol

The effect of autoclaving on the deprotenization process of shrimp shells was studied. The experimental conditions of the deproteinization process are shown in Table 3. The sugar solution was prepared by dissolving 20g galactose in I L deionized distilled water and then autoclaving (Model No. STM-E, Market Forge Sterilmatic, New York, USA) at 121˚C and 103.4 kPa for 30 min. Each reactor was loaded up to 75% of its capacity (200 g shells based on dry weight). An inoculum concentration of 1 × 107 spores per 1g shrimp shell waste was used. The initial moisture content of the shrimp shells was adjusted to 60% with the addition of sugar and spores solutions and the material was mixed thoroughly. Air was introduced inside each reactor at a flow rate of 5 VWM (I m air per g shells per minute). The experiment ran for 6 days. At the start of the experiment, the reactors were rotated (1 rpm) continuously for 30 min and then intermittently for 15 min every hour.


Experimental Analyses

The particle size distribution, moisture content, pH, galactose concentration, ammonium nitrogen, total Kjeldahl nitrogen, protein and chitin measurements were performed on the shrimp shells. During the course of the deproteinization process, shrimp shell samples of 10 g each were collected every 12 h and analyzed for moisture content, pH, galactose concentration, protease activity, ammonium nitrogen content, total Kjeldahl nitrogen content and protein content. Exhaust gas samples were also taken every 12 h and analyzed for carbon dioxide concentration. The bulk temperature was monitored and recorded every 10 minutes. The deproteinized shells were analyzed for chitin and appearance.


Particle Size Distribution

A known weight of shrimp shells were sieved for 30 min using a sieve shaker (Model RX-86, Fisher Scientific, Montreal, Quebec, Canada) with 7 different sieve sizes (6.300, 4.000, 2.000, 0.850, 0.300, 0.180, 0.075 mm aperture size). Each particle size fraction obtained was weighed and the percentage from the total weight was calculated.  Table 2 show the particle size distribution of the shrimp shells used in this study.


Moisture Contents

A sample of a known weight of shrimp shells was placed in a preweighed aluminum dish. The dish and content were weighed and then placed in an oven (Isotemp Oven, Model 655F, Fisher Scientific, Montreal, Quebec, Canada) at 105 ºC  for  24  hours.  The  aluminum  dish  along with the dried sample were first placed in a desiccator to cool down and then weighed. The moisture content was determined as follows:

                               (1)

Where:

MC   is the moisture content (%)

Wws   is the weight of the wet sample (g)

Wds    is the weight of the dry sample (g)   


pH

20 mL of deionized distilled water was added to one gram sample of shrimp shells and kept at room temperature (24 °C) for 30 min with frequent stirring using a stir plate (Thermix® Stirrer Model 120MR, Fisher Scientific, Montreal, Quebec, Canada). The sample was let stand for two minutes and the pH was then measured using a pH meter (Model 805MP, Fisher Scientific, Montreal, Quebec, Canada). 

Samples were washed thoroughly several times with deionized distilled water until the wash water was clear and then dried in an oven (Isotemp Oven, Model 655F, Fisher Scientific, Montreal, Quebec, Canada) at 60 ºC till constant weight. The dried shells were ground using a small conventional grinder (Hamilton Beach, Markham, Ontario, Canada). Ammonium nitrogen (NH4-N) of dry ground samples was determined directly using Kjeldahl system (KJELTEC AUTO 1030 Analyzer, Fisher Scientific, Montreal, Quebec, Canada).


Ammonium Nitrogen

Samples were washed thoroughly several times with deionized distilled water until the wash water was clear and then dried in an oven (Isotemp Oven, Model 655F, Fisher Scientific, Montreal, Quebec, Canada) at 60 ºC till constant weight. The dried shells were ground using a small conventional grinder (Hamilton Beach, Markham, Ontario, Canada). Ammonium nitrogen (NH4-N) of dry ground samples was determined directly using Kjeldahl system (KJELTEC AUTO 1030 Analyzer, Fisher Scientific, Montreal, Quebec, Canada).


Total Kjeldahl Nitrogen

Samples were washed thoroughly several times with deionized distilled water until the wash water was clear and then dried in an oven (Isotemp Oven, Model 655F, Fisher Scientific, Montreal, Quebec, Canada) at 60 ºC till constant weight. The dried shells were ground using a small conventional grinder (Hamilton Beach, Markham, Ontario, Canada) and then digested by heating the sample with concentrated sulfuric acid and kjeltabs (which contained 3.5 g K2SO4 and 0.0035 Se) for 45 min. The K2SO4 promotes the oxidation of organic matter by increasing the temperature of the digest (420 ºC) that resulted in the conversion of organic nitrogen to ammonium nitrogen. Se is a catalyst which increases the rate of oxidation of organic matter by sulfuric acid. 5 mL sulfuric acid with 1 kjeltab per 0.2 g dry weight sample were used in this study.  The total Kjeldahl nitrogen (TKN) of the digested samples was as determined using Kjeldahl system (KJELTEC AUTO 1030 Analyzer, Fisher Scientific, Montreal, Quebec, Canada).


Protein

The protein content of the samples was determined using the following equations which are based on the fact that the protein contains about 16% nitrogen:

(Org.-N)s = TKNs – (NH4-N)s                                              (2)

PRc = [(Org.-N)s – (Org.-N)c] × 6.25        (3)

Where:

PRc             is protein content (mg/kg)

(Org.-N)c    is organic nitrogen of the recovered chitin (mg/kg)

(Org.-N)s     is organic nitrogen of the sample (mg/kg)

TKNs          is total Kjeldahl nitrogen of the sample (mg/kg)

(NH4-N)s    is ammonium nitrogen of the sample (mg/kg)


Galactose Concentration

20 mL of deionized distilled water was added to one gram sample of fermented shrimp shells and kept at room temperature (24 °C) for 30 min with frequent stirring using a stir plate (Thermix® Stirrer Model 120MR, Fisher Scientific, Montreal, Quebec, Canada). The extract was then filtered under suction using coarse porosity filter paper (Reeve Angel Grade 202, Whitman Inc., Clifton, New Jersey, USA) and the supernatant was used for galactose concentration measurements using the phenol-sulfuric acid method which is based on the fact that simple sugars give a stable orange-yellow color when reacting with phenol and concentrated sulfuric acid [27]. The intensity of the color is proportional to the amount of galactose present in the sample and can be determined by measuring the absorbance at 492 nm.


A standard curve was developed from solutions of galactose and deionized distilled water with different concentrations. First, a standard solution of 100 µg/mL galactose was prepared by dissolving 10 mg galactose in 100 mL deionized distilled water. Then, a set of 6 solutions with galactose concentration of 5, 10, 15, 20, 25, and 30 µg/mL) were prepared. Finally, the absorbance was measured using a microplate reader (µQuant, Bio-Tek Instruments, Inc., Winooski, Vermont, USA) at 492 nm. A blank sample of pure deionized distilled water was used. The measured absorbances were plotted against the known galactose concentrations (µg/mL) as shown in Figure 4. The following linear relationship between the galactose concentration and the absorbance was obtained (R2= 0.98):

Cga = 416.67 (Ā492)                                     (4)

Where:

Cga    is the galactose concentration (µg/mL)


To measure galactose in the sample, 2 mL of each sample were transferred to a test tube and 1 mL of phenol solution and 5 mL of concentrated sulfuric acid (95-98%) were added to the tube. The tubes were then tight capped and the contents were mixed using a vortex mixer (Sybron Maxi Mix model M-16715, Thermolyne Corporation, Dubuque, Iowa, USA). The tubes were allowed to stand for 10 minutes at room temperature and then the contents were mixed again using the vortex mixer. The tubes were placed in a water bath (2850 Series, Fisher Scientific, Toronto, Ontario, Canada) at 30 ºC for 15 minutes after which the contents were mixed again using the vortex mixer. The tubes were allowed to stand for 30 minutes at room temperature. 200µL of each tube were carefully loaded into duplicate wells in a microliter plate and the absorbance was measured using a microplate reader (µQuant, Bio-Tek Instruments, Inc., Winooski, Vermont, USA). 


Protease Activity

Protease produced by A. niger was first extracted from the shrimp samples (1 g each) using 20 mL deionized distilled water and kept at room temperature for 30 min with continuous stirring using a stir plate (Thermix® Stirrer Model  120MR,  Fisher  Scientific,  Montreal, Quebec, Canada). The extract was then filtered under suction using coarse porosity filter paper (Reeve Angel Grade 202, Whitman Inc., Clifton, New Jersey, USA) and the supernatant was used for the assay of enzyme. Protease activity was measured using Protease Colorimetric Detection Kit (Product Code PC0100, Sigma, Saint Louis, Missouri, USA). The assay was based on using a casein substrate, which is cleaved by the protease to trichloroacetic acid soluble peptides. The formed peptides contain tyrosine and tryptophan residues, which react with the Folin and Ciocalteu’s reagent causing color change, which can be estimated calorimetrically at 660 nm using a microplate reader (µQuant, Bio-Tek Instruments, Inc., Winooski, Vermont, USA).


Chitin

The chitin content was determined based on the fact that chitin contains about 6.89% organic nitrogen [28]. In order to determine the chitin nitrogen, samples were first deproteinized and demineralized.


The deproteinization process was performed using 5% (w/v) NaOH solution. One gram of ground shrimp shell sample (dry weight) along with 100 mL of NaOH solution were placed in a 250 mL wide-mouth flask and the flask was covered with a piece of tin foil and sealed with a rubber band to ensure the retention of all reacted materials. The flask was then placed in a boiling water bath (2850 Series, Fisher Scientific, Toronto, Ontario, Canada) for 1 h. The sample was filtered under suction through a Buchner funnel with coarse porosity filter paper (Reeve Angel Grade 202, Whitman Inc., Clifton, New Jersey, USA) and washed thoroughly with deionized distilled water. The deproteinized sample was dried in an oven (Isotemp Oven, Model 655F, Fisher Scientific, Montreal, Quebec, Canada) at 60 ºC till constant weight. The weight of the recovered dry deproteinized sample was determined.


The deproteinized sample along with 50 mL of 1.0 M HCl were placed in a 250 mL wide-mouth flask and the flask was covered with a piece of tin foil and sealed with a rubber band to ensure the retention of all reacted materials. The flask was then placed in a boiling water bath for 1 h. The demineralized sample was then filtered under suction through a Buchner funnel with coarse porosity filter paper (Reeve Angel Grade 202, Whitman Inc., Clifton, New Jersey, USA) and washed thoroughly with deionized distilled water. The deproteinized-demineralized sample was then dried in an oven (Isotemp Oven, Model 655F, Fisher Scientific, Montreal, Quebec, Canada) at 60 ºC till constant weight. The weight of the recovered dry deproteinized-demineralized sample was determined. The ammonium and total kejldhal nitrogen analyses were performed on the dry deproteinized-demineralized sample and the chitin content was then calculated as follows:

(Org.-N)c = [(TKNc – (NH4-N)c] × Wc/Ws                  (5)

CHc = (Org.-N)c × 14.51                            (6)

Where:

CHc          is chitin content (mg/kg)

(Org.-N)c  is organic nitrogen of the recovered chitin

TKNc        is total Kjeldahl nitrogen of the recovered chitin (mg/kg)

(NH4-N)c  is ammonium nitrogen of the recovered chitin (mg/kg)

Wc             is weight of recovered chitin (g)

Ws             is weight of sample (g) 


Ash Content

The dried shrimp shells were analyzed for their ash content. A known weight of the material was placed in a preweighed aluminum dish. The dish and content were weighed and then placed in a muffle furnace (Isotemp® Muffle Furnace model 186A, Fisher Scientific, Montreal, Quebec, Canada) at 700 ºC for 2 hours. The dish with the content was taken from the muffle furnace, placed in a desiccator to cool down and then weighed. The ash content was determined as follows:   

                           (7)

Where:

AC    is the ash content (%)

Wa     is the weight of the ash (g)


Minerals

The dried shrimp shells were analyzed for their minerals content. Quantitative trace element analyses (magnesium, calcium, manganese, potassium, sodium, iron, silicon, aluminum, titanium and copper) were performed on the ash using an Atomic Absorption Spectrophotometer (SpectrAA 55B, Varion, Mulgrave, Victoria, Australia) in the Minerals Engineering Center, Dalhousie University, Halifax, Nova Scotia. For magnesium, calcium, manganese, potassium, sodium, iron and copper analyses, the samples were first digested with hydrochloric, nitric, hydrofluoric and perchloric acids (30, 10, 10 and 5 mL/g sample, respectively) in a closed vessel at a temperature of 100 ºC and then the elements were determined by flame atomic absorption with detection limit of 1 ppm. For silicon, aluminum and titanium analyses, 1 g of the sample was fused with a flux of lithium metaborate and lithium tetraborate and leached with 1:9 nitric acid. Sulfur was determined with Leco Sulfur analyzer along with Leco Induction Furnace (Leco Corporation, St. Joseph, Michigan, USA). Phosphorus was determined as P2O5 by a colorimetric method using spectrophotometer with micro flow-thru system (Spectoronic 100, Bausch & Lomb Incorporation, Rochester, New York, USA) at 430 nm.


Visualization of Shrimp Shells

The shrimp shells were visually inspected at the end of the deproteinization with the naked eye as well as under the incident light stereomicroscope (Carl Zeiss Stemi SV8, Carl Zeiss Canada Ltd., Toronto, Ontario, Canada) at a magnification of 60X. The stereoscope was equipped with a cold light source (SCOHTT KL 1500, SCHOTT North America Inc., New York, USA) and a single chip CCD color video camera (Sony DXC-101, Sony of Canada Ltd., Toronto, Ontario, Canada).


RESULTS AND DISCUSSION

The effect of autoclaving on various deproteinization parameters (temperature, pH, moisture content, galactose utilization, carbon dioxide evolution, protease activity, residual protein and chitin concentration) of ground shrimp shells supplemented with galactose as a carbon source was investigated. 


Temperature

Figure 5 shows the changes in the temperature of the shrimp shells and the exhaust gas during the course of deproteinization as affected by autoclaving. The values are the average of three replicates. The coefficient of variation ranged from 0.40 to 4.32. 

The solid state fermentation process is an exothermic reaction in nature generating heat that gives rise to the temperature of the medium [29].  The microorganisms utilize organic carbon and micronutrients for synthesis of new microbial cells, product formation and energy generation [20,30]. The heat stored in the bioreactor is the net of metabolic heat production minus the heat losses (by conduction through the shrimp shells and the bioreactor walls, by convection with the exhaust gas and latent heat through water evaporation from the shrimp shells).


In this study, the average inlet air temperature and initial temperature of the shrimp shells were 23.3 ± 0.54 °C and 21.8 ± 0.34 °C, respectively. The temperatures of the shrimp shells and the exhaust gas declined at the beginning of the fermentation process (lag period) as the heat losses from the bioreactor were higher than the heat generated in the bioreactor by microorganisms. After 12h, the temperature of the shrimp shells started to rise as the heat generation by metabolic activity exceeded the heat losses. The rise in the exhaust gas temperature was due to heat losses from the shrimp shells by convection. The temperatures of the shrimp shells and exhaust gas reached maximum values of 37.5 and 33.6 ºC and 28.9 and 27.9 ºC after 60 and 72 h for the autoclaved and unautoclaved shrimp shells, respectively. 


Ghildyal et al. [31] and Pandey [32] reported that temperatures in the middle of the bed can reach about 20 ˚C higher than the temperature of the inlet air.  Saucedo-Castañeda et al. [29] reported an axial temperature gradient of 0.17 ˚C/cm and a radial temperature gradient of 5 ˚C/cm in the bioreactor during the fermentation of cassava using A. niger. In this study, the peak temperature of the shrimp shells material was about 10.3-14.6 ˚C higher than that of the inlet air temperature and there were no temperature gradients in the radial or axial direction because of mixing.


Ben-Hassan et al. [33] and Ghaly et al. [34] showed that the microbial growth and the temperature curves are similar in shape and the temperature curve can be used to determine

microbial kinetics. The growth kinetics (Table 4) was determined from the temperature data for the autoclaved and unautoclaved shrimp shells according to the procedures described by Ghaly et al. [35]. Lag periods of 12 and 12 h were observed for the unautoclaved and autoclaved shells, respectively. The exponential growth phase was 48 h for autoclaved shells and 24 h for theunautoclaved shells. The stationary phase was 4 h for autoclaved shells and 23 h for unautoclaved shells. The maximum temperatures (37.5 and 33.0 °C) were attained after 60 and 48 h for autoclaved and unautoclaved shells, respectively. The specific growth rate for A. niger was 0.022 h-1 for autoclaved shells and 0.018 h-1 for unautoclaved shells.


pH

Figure 6 shows the change in the pH of shrimp shells during the course of deproteinization as affected by initial autoclaving of the shrimp shells. The values are the average of three replicates. The coefficient of variation ranged from 0.46 to 3.51%.


For the run with autoclaved shrimp shells, the initial pH of 8.64 decreased with time reaching 7.34 after 54 h, then increased reaching 8.16 at the 84 h and finally declined to 7.66 by the end of the deproteinization process. For the run with unautoclaved shrimp shells, the initial pH of 8.64 decreased with time reaching 7.05 after 60 h, then increased reaching 7.35 at the 108 h and finally declined to 7.16 by the end of the deproteinization process. Villegas et al. [36] stated that changes in pH during fermentation are linked to the ionic balance established by substrate uptake and product formation. The decrease in the pH of the shrimp shells observed in this study was due to the production of acid protease while the increase in the pH of the shrimp shell was due to the buffering capacity of the calcium carbonate released from the shrimp shells [37] as well as the production of ammonium nitrogen as reported by Yang and Lin [37]. 

Zakaria et al.[2] reported a drop in the pH to a value of 5 over the first 48 h of lactic acid fermentation of scampi waste after which the pH increased reaching a final value of 6.6 as a result of the buffering capacity of the solubilized calcium. Beaney et al. [12] reported rapid decrease in pH to 3.5 over 7 days during lactic acid fermentation of prawn shells as a result of metabolic lactic acid production. Beaney et al. [12] reported rapid decrease in pH to 3.5 over 7 days during lactic acid fermentation of prawn shells as a result of metabolic   lactic   acid   production.   Andrade  et  al.  [38] reported a decrease in the medium pH during the period of protease production using Mucor circinelloides as a result of metabolites accumulation resulting from D-glucose degradation.


Moisture Content

Figure 7 shows the change in moisture content of the shrimp shells during the course of deproteinization as affected by the initial autoclaving of the shrimp shells. The values are the average of two replicates. The coefficient of variation ranged from 0.16 to 6.22%. 

The net moisture content of the shrimp shell bed can be defined as follows:

MCnet = MCi + MCm – MCe                                                (7)

Where:

MCnet    is the net moisture content (%)

MCi          is the initial moisture content (%)

MCm     is the metabolic moisture content (%)

MCe      is the moisture lost through evaporation with the exhaust gas (%)


The results showed that the moisture content profiles of the autoclaved and unautoclaved shrimp shells were similar. The moisture content of the shrimp shells declined first slowly in the first 37 h from the initial value of 60% to 54.8%, then sharply reaching values of 23.2% and 35.1% by the 72 h and finally decreased slowly reaching final values of 25.54 and 26.72% at the end of the deproteinization process for the autoclaved and unautoclaved shrimp shells, respectively. The initial slow decrease in the moisture content was due to the initial low temperature which affected the amount of water lost by evaporation while the steep decline in the moisture content observed thereafter indicated higher loss of that moisture as the water lost by evaporation was higher than that produced by metabolic activities.


The optimum moisture content for solid state fermentation is in the range of 40-60%. High moisture content (above 60%) cause packing of the substrate and prevention of gas exchange and low moisture content (below 40%) cause the substrate to be too dry for microbial growth and product formation [37]. Yang and Chiu [39] reported optimum initial moisture content between 50-58% for the production of protease by solid-state fermentation. Diaz et al. [40] reported that microbial activity are inhibited at moisture contents below 40% and completely cease at moisture contents below 12%. In this study, the initial moisture content of the shrimp shells was adjusted to 60% which fell below 40% after 60 h and reached 25.54 and 26.72% at the end of the deproteinization process for the autoclaved and unautoclaved shrimp shells, respectively. The low moisture content observed in this study may have negatively affected fungus growth and protease production and in turn the deproteinization process. To maintain the moisture of the substrate inside the bioreactor at the desired level, the exhaust gas should be passed through a condensation tower and the recovered water pumped back into the bioreactor through the aeration tube.


Galactose Utilization

Figure 8 shows the changes in residual galactose concentration in the shrimp shells during the course of deproteinization as affected by the initial autoclaving of the shrimp shells. The values are the average of three replicates. The coefficient of variation ranged from 1.92 to 14.51%.

The optimum galactose concentration of 20% w/w was used in this study as recommended by Mahmoud and Ghaly [41]. The results showed that the rate of galactose utilization was slightly higher for autoclaved shrimp shells comparedto that of unautoclaved shrimp shells. The galactose concentration decreased gradually over the course of the deproteinization process reaching about 1.48 and 2.67% by the end of the deproteinization process for the autoclaved and unautoclaved shells, respectively.


Carbon Dioxide Evolution

Figure 9 shows the effect of autoclaving of shrimp shells on carbon dioxide concentration in the exhaust gas. The values are the average of three replicates. The coefficient of variation ranged from 2.01 to 10.98%. The results showed that carbon dioxide concentrations in the exhaust gas measured during the deproteinization process for both the autoclaved and unautoclaved shells had similar profiles. However, higher concentrations of CO2 were observed with the autoclaved shrimp shells. The carbon dioxide increased with time reaching its peak of 0.37% and 0.32% at 60 h and then declined reaching 0.06 and 0.06% by the end of the experiment for the runs with autoclave and unautoclaved shrimp shells, respectively. 

Temperature and carbon dioxide evolution are considered strong indicators of microbial activity during solid-state fermentation [42,43]. Carrizalez et al. [44] used carbon dioxide measurements for the determination of microbial growth of A. niger culture on cassava flour. Rathbun and Shuler [45] stated that the relationship of CO2 evolution and microbial growth is accurate and used the rate of COevolution (gas flow time mole fraction CO2) as a measure of the rate of microbial growth. In this study, The CO2 concentration also followed the same trend as the temperature of the shrimp shells. A strong correlation (R2 = 0.9) between the concentration of carbon dioxide in the exhaust gas and the temperature of the shrimp shells was obtained (Figure 10).


Protease Activity

Figure 11 shows the effect of autoclaving of shrimp shells on protease activity during the course of deproteinization. The values are presented in units per gram dry shrimp shells and the values are the average of three replicates. The coefficient of variation ranged from 2.20 to 6.98%.


The results obtained from this study revealed the ability of A. niger to produce extracellular proteases that resulted in deprorinization of the protein in the shrimp shells. Fish proteins are complex molecules consisting of chains of amino acids linked together by peptide bonds. Proteases are proteins structured in such a way that allow them to act as catalysts in the breakage of peptide bonds through a process called hydrolysis according to the following equation:


      (8)


Bustos and Healy [8] stated that the degree to which a microorganism will hydrolyze a protein substrate depends on its capacity to produce the required protease and the stability of a such protease under the reaction conditions. The results obtained from this study showed that the protease activity was higher in case of autoclaved shrimp shells. The protease activity increased with time from an initial value of 0.67 unit/g dry shrimp shell to final values of 4.21 and 2.44 unit/g dry shrimp shell after 5 d for the autoclaved and unautoclaved shrimp shells, respectively. Ashour et al. [46] reported that protease yield from the fungus A. niger during cheese whey fermentation  increased with incubation period and reached a maximum value after 6 days. Teng et al. [19] reported protease activities in the range of 0.8-6.8 units (one unit activity was defined as 1 µM of tyrosine produced in 1 min) for 17 A. niger strains after 5 days of incubation of the spores.


Protein Concentration

Figure 12 shows the effect of autoclaving of shrimp shells on chitin content of the shrimp shells during the course of deproteinization. The values are presented based on the dry weight of the samples and are the average of three replicates. The coefficient of variation ranged from 1.22 to 6.18%.

The results showed that protein concentration of the shrimp shells decreased with the deproteinization time as a result of protein break down by the proteolytic enzymes produced by A. niger. However, the protein concentration in the shrimp shells was lower in case of autoclaved shrimp shells. The protein concentration decreased from the initial value of 30.84 to final values of 20.59% and 22.94% by the end of the deproteinization process for the autoclaved and unautoclaved shells, respectively. Zakaria et al. [2] used lactic acid fermentation for chitin recovery from scampi waste and reported 77.5% deproteinization efficiency after 5 days. Beaney et al. [12] reported 50% decrease of the original protein concentration in prawn shell waste using lactic acid fermentation and stated that complete deproteinization through a purely biotechnological process seems hard to achieve.

The low deproteinization efficiency observed in this study could be due to the high pH of the shrimp shells that might have interfered with protease synthesis and/or activity. Diniz and Martin [47] stated that the extent of hydrolytic degradation of protein depends on pH, temperature, extend of native protein denaturation, concentration and specificity of the enzyme, composition and the molecular weight distribution of the peptides in the protein and presence of inhibitory substances. The optimum pH of acid protease production from A. niger lies in the range of 2.0 - 3.0 and the pH of the shrimp shells during the entire deproteinization process was far (7.05-8.64) from being optimum for enzymes production.

Chitin Concentration

Figure 13 shows the effect of autoclaving of shrimp shells on residual protein in the shrimp shells during the course of deproteinization. The values are presented based on the dry weight of the samples and are the average of three replicates. The coefficient of variation ranged from 2.32 to 4.48%.

The chitin concentration was determined in the deproteinized samples without demineralization. The shrimp shells used in this study contained31.73% minerals.  The results showed that the chitin concentration in the shells increased from an initial value of 16.56 % to final values of 22.68% and 20.93% over the course of the deproteinization process for the autoclaved and unautoclaved shrimp shells, respectively. Zakaria et al. [2] reported an increase in the concentration of chitin from 12.05% to 17.48% as a result of lactic acid fermentation of scampi waste. Cira et al. [48] reported increases in chitin concentration from 11.4-13.1% to 20.3-23.2% as a result of lactic acid fermentation of shrimp waste.

Visualization of Shrimp Shells

Figure 14 shows the visual appearance of autoclaved and unautoclaved shrimp shells at the end of the deproteinization process (120 h). Figure 15 shows the appearance of autoclaved and unautoclaved shrimp shells under the microscope (60X magnification) at the end of the deproteinization process. Samples of the autoclaved and unautoclaved shrimp shells were plated on potato dextrose agar after 24 and 72 h of deproteinization. Figure 16 shows the effect of autoclaving on the microbial population after 24 and 72 h fermentation time.

The spent shrimp shells obtained from both runs had a pale pink-orange color with some tan patches. More of the white precipitant accumulated on the surface of the autoclaved shrimp shells compared to the one that was accumulated on the surface of the unautoclaved shrimp shells as noticed under the microscope (60X magnification). The existence of the pink-orange color was an indication of the presence of pigments, which were not utilized during the fermentation process. The extent of shrimp shell agglomeration was much higher in case of unautoclaved shrimp shells than in the case of autoclaved shrimp shells. The plates showed the existence of unidentified microorganisms on both autoclaved and unautoclaved shells. However, the population of the unidentified microorganisms was much higher in the case of unautoclaved shells.

CONCLUSIONS

The results obtained from this study revealed the ability of A. niger to produce extracellular proteases in the presence of shrimp shell protein and galactose as carbon source. As a result of proteases production, the shrimp shells were deproteinized to a certain degree by the end of the deproteinization process.

The temperatures of the shrimp shells and the exhaust gas declined at the beginning of the deproteinization process (lag period) as the heat losses from the bioreactor were higher than the heat generated by metabolic activities. Once the exponential growth phase started, the temperatures of the shrimp shells and exhaust gas increased and reached maximum values of 37.5 and 33.6 ºC and 28.9 and 27.9 ºC after 60 and 72 h for the autoclaved and unautoclaved shrimp shells, respectively. The peak temperature of the shrimp shells in the bioreactor was about 10.3-14.6 ˚C higher than that of the inlet air temperature and there were no temperature gradients in the radial or axial direction because of mixing. Since the optimum temperature for protease activity is in the range of 20-26 ˚C, removal of metabolic heat during the deproteinization process might have a positive effect on the growth of A. niger and the production of acid protease.

The carbon dioxide increased with time reaching its peak of 0.37% and 0.32% at 60 h and then declined reaching 0.06 and 0.06% by the end of the experiment for the runs with autoclave and unautoclaved shrimp shells, respectively.  The CO2 concentration followed a similar trend to that of the temperature of the shrimp shells and a strong correlation between the concentration of carbon dioxide in the exhaust gas and the temperature of the shrimp shells was obtained.   

The initial pH of 8.64 decreased first with time reaching 7.34 after 54 h and 7.05 after 60 h and then increased reaching 8.16 after 84 h and 7.35 after 108 h and finally declined to 7.66 and 7.17 by the end of the deproteinization process for the autoclaved and ubautoclaved shrimp shells, respectively. The decrease in the pH of the shrimp shells was due to the production of acid protease while the increase in the pH of the shrimp shell was due to the buffering capacity of the calcium carbonate released from the shrimp shells and the production of ammonium nitrogen.  

An initial moisture content of 60% was used in this study which is ideal for bacterial growth while moisture content of 45% is adequate for fungal growth. Thus, the high initial moisture content may have enhanced the growth of contaminant microbes in the unautoclaved shells at the beginning of the experiment (lag period). However, the temperature increased during the exponential growth phase as a result of the metabolic heat produced by A. niger resulting in a significant reduction in the moisture content The moisture content of the shrimp shells fell below 40% after 60 h and reached 25.54-26.72% at the end of the deproteinization process. The low moisture content may have affected the growth and metabolic activity of A, niger. The recovery of the water lost from the bioreactor with the exhaust gas is recommended. The exhaust gas can be passed through a condensation tower and the recovered water can be pumped back into the bioreactor through the aeration tube.

The galactose concentration decreased gradually over the course of the deproteinization process reaching about 1.48 and 2.67% by the end of the deproteinization process for the autoclaved and unautoclaved shells, respectively. Since most of the galactose was utilized after 60 h, a fed-batch galactose addition would result in longer exponential growth phase and more acid protease production and consequently more protein removal.  

The protein concentration decreased from the initial value of 30.84 to final values of 20.59% and 22.94% by the end of the deproteinization process for the autoclaved and unautoclaved shells, respectively. The deproteinization efficiencies of autoclaved and unautoclaved shells were 33.23% and 25.62% for, respectively. The reduction in the protein concentration at the end of the deproteinization process did not correspond to the increase in the protease activity. The low deproteinization efficiency observed in this study could be due to the high initial pH of the shrimp shells and the high temperature in the bioreactor which might have interfered with protease synthesis and/or activity. The optimum pH of acid protease from A. niger lies in the range of 2.0 - 3.0 and the pH of the shrimp shells during the entire deproteinization process was far (7.05-8.64) from being optimum for enzymes production. The low deproteinization attained with unautoclaved shells might be due to the growth of the indigenous microorganisms in the shrimp shells which might affect the growth and metabolic activity of A. niger. Also, denaturation of proteins due to exposure to high temperatures make them more susceptible to enzymatic hydrolysis.

The protease activity of the autoclave shells was significantly higher (1.7 fold) than that of the unautoclaved shells. The protease activity increased with time from an initial value of 0.67 unit/g dry shrimp shell to final values of 4.21 and 2.44 unit/g dry shrimp shell for the autoclaved and unautoclaved shrimp shells, respectively.

The shrimp shells used in this study contained 31.73% minerals and the chitin concentration were determined in the deproteinized samples without demineralization. It increased from an initial value of 16.56 to final values of 22.68% and 20.93% over the course of the deproteinization process for the autoclaved and unautoclaved shrimp shells, respectively.

The spent shrimp shells obtained from both runs had a pale pink-orange color with some tan patches. More of the white precipitant accumulated on the surface of the autoclaved shrimp shells compared to that accumulated on the surface of the unautoclaved shrimp shells. The pink-orange color was an indication of the presence of pigments which were not utilized during the fermentation process. The extent of shrimp shell agglomeration was much higher in the case of unautoclaved shrimp shells compared to the autoclaved shrimp shells. Unidentified microorganisms were found in both the autoclaved and unautoclaved shells with larger populations in the case of unautoclaved shells.

ACKNOWLEDGMENTS

The research was funded by the National science and Engineering Council (NSERC) of Canada. The support of Dalhousie University and Cairo University is highly appreciated.

1. Hall GM,  Reid CL, Zakaria Z (1994) Fermentation of prawn waste by lactic acid bacteria: Sea Fisheries Institute Gdynia, Poland. Proceeding from the 6th International Conference on Chitin and Chitosan, Wirtschaftsverlag, NW. 

2.     Zakaria Z, Hall GM, Shama S (1998) Lactic acid fermentation of scampi waste in a rotating horizontal bioreactor for chitin recovery. Process Biochemistry 33: 1-6. doi:10.1016/S0032-9592(97)00069-1

3.     Hansen HØ, Aschan  M (2000) Growth size- and age-at-maturity of shrimp, Pandalus Borealis, at Svalbard related to environmental parameters. Journal of Northwest Atlantic Fishery Science 27: 83-91. doi:10.2960/J.v27.a8

4.     Shahidi F, Synowiecki J (1992) Quality and compositional characteristics of Newfoundland shellfish processing discards: Advances in Chitin and Chitosan, Brine. Elsevier Applied Science, England. ISBN:1566765730

5.     DFO (2014) Northeren Shrimp Fisheries. Department of Fisheries and Oceans, Ottawa, Ontario, Canada. http://www.dfo-mpo.gc.ca/index-eng.htm

6.     Zakaria MB, Jais MJ, Alimuniar A, Harahap ZA, Nagah WSW (1995) Chitosan as a chemical agent in the treatment of water and waste waters: Chitin and Chitosan the Versatile Environmentally Friendly Modern Materials. Universiti Kebangsaan Malaysia, Malaysia.

7.     Dutta PK, Dutta J, Tripathi VS (2001) Chitin and chitosan: chemistry, properties and applications. Journal of Scientific and Industrial Research 63: 20-31. ISSN:0975-1084

8.     Bustos RO, Healy MG (1994) Microbial deproteinisation of waste prawn shell. Institution of Chemical Engineers Symposium Series, Institution of Chemical Engineers, Rugby, England.

9.     Ghanem A, Ghaly AE, Chaulk M (2003) Effect of shrimp processing procedures on the quality and quantity of extracted chitin from the shells of northern shrimp Pandalus borealis. Journal of Aquatic Food Product Technology 12: 63-79. doi: 10.1300/J030v12n04_05

10.  Hall GM, De Silva S (1992) Lactic acid fermentation of shrimp (Peneaus monodon) waste for chitin recovery: Advances in Chitin and Chitosan, Brine. Elsevier Applied Sciences, London, UK. doi: 10.1002/pi.4990310419

11.  Hall GM, Reid CL (1995) Scale up of lactic acid fermentation process for the recovery of chitin from prawn waste: Chitin and Chitosan the Versatile Environmentally Friendly Modern Materials. Universiti Kebangsaan Malaysia.  ISBN-13:9789679423037

12.  Beaney P, Mendoza JL, Healy M (2005) Comparison of chitins produced by chemical and bioprocessing methods. Journal of Chemical Technology and Biotechnology 80: 145-150. doi: 10.1002/jctb.1164

13.  Shimahara K, Ohkouchi K, Ikeda M (1982) A new isolation method of crustacean chitin using a proteolytic bacterium, Pseudomonas maltophilia: Chitin and Chitosan, Proceeding of the second International Conference on Chitin and Chitosan. Japanese Society of Chitin and Chitosan.

14.  Cano-Lopez A, Simpson BK, Haard NF (1987) Extraction of carotenoprotein from shrimp process wastes with the aid of trypsin from Atlantic cod. Journal of Food Science 52: 503-506. doi: 10.1111/j.1365-2621.1987.tb06656.x

15.  Shimahara K, Takiguchi Y (1988) Preparation of crustacean chitin. Methods in enzymology 417-423. doi: 10.1016/0076-6879(88)61049-4

16.  Gagné N, Simpson BK (1993) Use of proteolytic enzymes to facilitate the recovery of chitin from shrimp wastes. Food Biotechnology 7: 253-263. doi: 10.1080/08905439309549861

17.  Bustos RO, Romo CR, Healy MG (1999) Purification of trypsin-like enzymes from Antarctic krill processing wastewater. Process Biochemistry 161: 327-333.  doi:10.1016/S0032-9592(99)00075-8

18.  Wang S, Chio S (1998) Deproteinization of shrimp and crab shell with the protease of Pseudomonas eruginosa K-187. Enzyme and Microbial Technology 22: 629-633. doi:10.1016/S0141-0229(97)00264-0

19.  Teng WL, Khor E, Tan TK, Lim LY, Tan SC (2001) Cocurrent production of chitin from shrimp shells and fungi. Carbohydrate Research 332: 305-316.  doi:10.1016/S0008-6215(01)00084-2

20.  Pandey A (1992) Recent process developments in solid-state fermentation. Process Biochemistry 27: 109-117. doi:10.1016/0032-9592(92)80017-W

21.  Berka, RM, Dunn-Coleman N, Ward M (1992) Industrial enzymes from Aspergillus species: Aspergillus Biology and Industrial Applications. Butterworth-Heinemann, Stoneham, MA. ISSN:0740-7378

22.  Ray MK, Devi KU, Kumar GS, Shivaji S (1992) Extracellular protease from Antarctic yeast Candida humicola. Applied and Environmental Microbiology 58: 1918-1923.  ISSN:0099-2240

23.  Naidu GSN, Panda T (2003) Studies on pH and thermal deactivation of pectolytic enzymes from Aspergillus niger. Biochemical Engineering Journal 16: 57-67.        doi:10.1016/S1369-703X(03)00022-6

24.  Kong N (1975) A feasibility study of new routes to the marine polymers chitin and chitosan. Unpublished master’s thesis. University of Washington, Washington, USA.

25.  Collins CH, Lyne PM (1984) Microbiological Methods. Butterworths & Co, Ltd. London, UK. ISBN:0408700483

26.  Wehr HM, Frank F (2004) Standard methods for the examination of dairy products.  American Public Health Association. ISBN: 978-0-87553-264-6

27.  Chow PS, Landhausser SM (2004) A Method for routine measurement of total sugars in woody plant tissues. Tree Physiology 24: 1129-1136. doi: 10.1093/treephys/24.10.1129

28.  Zakaria Z (1997) Lactic acid purification of chitin from prawn waste using a horizontal rotating bioreactor. Unpublished PhD thesis, Loughborough University. Loughborough, UK. doi: 10.1016/S0032-9592(97)00069-1

29.  Saucedo-Castañeda G, Gutiérrez-Rojas M, Bacquet G, Raimbault M, Viniegra-González G (1990) Heat transfer simulation in solid substrate fermentation. Biotechnology and Bioengineering 35: 802-808. doi: 10.1002/bit.260350808

30.  Ghildyal NP, Ramakrishna M, Lonsane BK, Karanth NG, Krishnaiah MM (1993) Temperature variations and amyloglucosidase levels at different bed depths in a solid state fermentation system. The Chemical Engineering Journal 51: B17-B23.        doi:10.1016/0300-9467(93)80019-K

31.  Ghildyal NP, Gowthaman MK, Raghava Rao KSMS, Karanth NG (1994) Interaction of transport resistances with biochemical reaction in packed-bed solid-state fermentors: effect of temperature gradients. Enzyme and Microbial Technology 16: 253-257.        doi:10.1016/0141-0229(94)90051-5

32.  Pandey A (2003) Solid-state fermentation. Biochemical Engineering Journal 13: 81-84. ISSN 1369-703X

33.  Ben-Hassan RM, Ghaly AE, Ben-Abdallah N (1993) Heat generation during batch and continuous production of single cell protein from cheese whey. Biomass and Bioenergy 4: 213-225. doi:10.1016/0961-9534(93)90060-H

34.  Ghaly AE, Kamal M, Avery A (2003) Influence of temperature rise on kineticparameters during propagation of Kluyveromyces fragilis in cheese whey under ambient condition. World Journal of Microbiology 19: 741-749. doi: 10.1023/A:1025148022934

35.  Ghaly AE, Kok R, Ingrahm JM (1989) Growth rate determination of heterogeneous microbial population in swine manure.  Applied Biochemistry and Biotechnology 22: 59-78. ISSN:0273-2289

36.  Villegas E, Aubague S, Alcantara L, Auria R, Revah S (1993) Solid state fermentation: acid protease production in controlled CO2 and O2 environments. Biotechnology Advances 11: 387-397. ISSN:0734-9750

37.  Yang FC, Lin IH (1998) Production of acid protease using thin stillage from a rice-spirit distillery by Aspergillus niger. Enzyme and Microbial Technology 23: 397-402.  doi:10.1016/S0141-0229(98)00070-2

38.  Andrade VS, Sarubbo LA, Fukushima K, Miyaji M, Nishimura K, et al. (2002) Production of extracellular proteases by Mucor circinelloides using D-Glucose as carbon source/substrate. Brazilian Journal of Microbiology 33: 106-110. http://dx.doi.org/10.1590/S1517-83822002000200002 

39.  Yang SS, Chui WF (1986) Protease production with sweet potato residue by solid state fermentation.  Zhonghua Minguo Weishengwu Ji Mianyixue Zazhi 19: 276-288.  ISSN:0253-2662

40.  Diaz, L. F., Savage, Eggerth G. M. and Golueke (1993) Composting and Recycling of Municipal Solid Waste. Lewis Publishers, Boca Raton, FL. ISBN: 0873715632

41.  Mahmoud NS, Ghaly AE (2015) Solid state deproteinization of shrimp shells by A. niger with galactose as a carbon supplement. American Journal of Biochemistry and Biotechnology. http://hdl.handle.net/10222/54765

42.  Lonsane BK, Ghildyal NP, Budiatman S, Ramakrishna SV (1985) ngineering aspects of solid state fermentation. Enzyme and Microbial Technology 7: 258-265.  doi:10.1016/0141-0229(85)90083-3

43.  Bellon-Maurel V, Orliac  O, Christen P (2003) Sensors and measurements in solid state fermentation: a review. Process Biochemistry 38: 881-896.  doi:10.1016/S0032-9592(02)00093-6

44.  Carrizalez V, Rodriguez  H, Sardina I (1981) Determination of the specific growth of molds on semisolid cultures. Biotechnology and Bioengineering 23: 321-333.  doi: 10.1002/bit.260230207

45.  Rathbun BL, Shuler ML (1983) Heat and mass transfer effects in static solid-substrate fermentation design of fermentation chambers. Biotechnology and Bioengineering 25: 929-938. doi: 10.1002/bit.260250405

46.  Ashour SA, El-Shora HM, Metwally M, Habib SA (1996) Fungal fermentation of whey incorporated with certain supplements for the production of proteases. Microbios 86: 59-69.  doi: 10.1007/BF03175548

47.  Diniz, FM, Martin AM (1997) Fish protein hydrolysates by enzymatic processing. Agro Food Industry Hi-Tech 8: 9-13.

48.  Cira LA, Huerta S, Hall GM, Shirai K (2002) Pilot scale lactic acid fermentation of shrimp wastes for chitin recovery. Process Biochem 37: 1359-1366.        doi:10.1016/S0032-9592(02)00008-0

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