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\title{Microalgae Growth in Qatar for CO2 Capture and Biodiesel Feedstock Production}
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\begin{document}

             \author[1]{Ihab H.  Farag}

             \author[2]{Dr. Rebecca J.  Wilson}

             \author[3]{Ghada  Salama}

             \affil[1]{  University of New Hampshire, Durham, USA.}

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\date{\small \em Received: 6 December 2011 Accepted: 31 December 2011 Published: 15 January 2012}

\maketitle


\begin{abstract}
        


Demands for and prices of liquid petroleum fuels are increasing. This challenge is motivating the development of alternative fuels, like biodiesel from non-food sources. Microalgae are a promising source of oil feedstock for biodiesel. Growing microalgae indoors uses water, chemical nutrients, artificial lights, and energy for harvesting, drying and oil extraction. The economics would be greatly improved if microalgae are grown outdoors in a hot sunny climate where the light energy is free and the temperature is adequate for growth. Using non-potable water (such as available and free salt-water) would reduce the water footprint. Open pond systems have low capital and operating costs and are wellsuited for growing microalgae in salty water. The ideal location for growing microalgae outdoors is a non-arable land that cannot be used for agriculture (such as Qatar desert). The purpose of this research is to study the growth of salt-water microalgae outdoors in Qatar?s hot sunny environment and compare it to indoor growth. Three Dunaliella microalgae (Bardawil, Parva and Salina) were grown in Persian Gulf saltwater medium. A fish tank photobioreactor was used to simulate an open pond. Dunaliella Bardawil provided the highest microalgae oil feedstock for biodiesel production, with a production rate of 20 mg dry algae/L-day, an oil content of 5.7 g oil/100 g dry algae, and oil production rate of 1.14 mg oil/L-day. The operation had a carbon sequestration efficiency of 6.5% and a photosynthetic efficiency of 1.11%. Among the algae tested, Dunaliella Bardawil is the optimal candidate for growth in Qatar conditions using an open pond system.

\end{abstract}


\keywords{Demands for and prices of liquid petroleum fuels are increasing. This challenge is motivating the development of alternative fuels, like biodiesel fro}

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\let\tabcellsep& 	 	 		 \par
are increasing. This challenge is motivating the development of alternative fuels, like biodiesel from non-food sources. Microalgae are a promising source of oil feedstock for biodiesel. Growing microalgae indoors uses water, chemical nutrients, artificial lights, and energy for harvesting, drying and oil extraction. The economics would be greatly improved if microalgae are grown outdoors in a hot sunny climate where the light energy is free and the temperature is adequate for growth. Using non-potable water (such as available and free salt-water) would reduce the water footprint. Open pond systems have low capital and operating costs and are wellsuited for growing microalgae in salty water. The ideal location for growing microalgae outdoors is a non-arable land that cannot be used for agriculture (such as Qatar desert). The purpose of this research is to study the growth of salt-water microalgae outdoors in Qatar's hot sunny environment and compare it to indoor growth. Three Dunaliella microalgae  {\ref (Bardawil, Parva and Salina)} were grown in Persian Gulf saltwater medium. A fish tank photobioreactor was used to simulate an open pond. Dunaliella Bardawil provided the highest microalgae oil feedstock for biodiesel production, with a production rate of 20 mg dry algae/L-day, an oil content of 5.7 g oil/100 g dry algae, and oil production rate of 1.14 mg oil/L-day. The operation had a carbon sequestration efficiency of 6.5\% and a photosynthetic efficiency of 1.11\%. Among the algae tested, Dunaliella Bardawil is the optimal candidate for growth in Qatar conditions using an open pond system.\par
iodiesel is a plant-derived biofuel intended to replace petroleum diesel. It is biodegradable, essentially CO 2 neutral, and much less toxic than petro diesel. It is made in a processor \hyperref[b28]{(Wilson and Farag, 2012)} by the transesterification of a triacylglycerides (TAGs)-containing oil feedstock (e.g., oils of soybean, rapeseed, maize and Jatropha, \hyperref[b30]{Tewfik et al., 2012)} and an alcohol (e.g., methanol or ethanol). One of the very promising oil feedstocks for biodiesel is microalgae oil (Nkongolo, 2010, Nkongolo and Farag, 2012, \hyperref[b10]{Chaput et al. 2012}\hyperref[b26]{, Zuka et al., 2012)}. 
\section[{B}]{B}\par
Microalgae are plant-like cells. They require a nutrient medium (water + nitrogen + phosphorous + other nutrients), CO 2 and light energy to do photosynthesis and grow. The simplest technique to grow microalgae in hot sunny areas like Qatar is in open ponds. During photosynthesis the algae capture the light energy and use it for carbon fixation, i.e., convert CO 2 (absorbed from air or water) to glucose and release oxygen. Up to 50\% is converted into TAG-containing lipids (oil) that can be used as a biodiesel feedstock in the transesterification reaction  {\ref (Scott, 2010)}. Table  {\ref 1} shows that microalgae, e.g., Nannochloropsis or Chlorella, have the potential to produce more oil feedstock for biodiesel than other crops. This is due to the simple cell structure and surface water interface. Algae cells have efficient and easy access to dissolved CO 2 and other nutrients while growing suspended in water. Hence algae are considered excellent CO 2 capture and use (CCU) systems. Estimates are that 2.5 tonnes of CO 2 are needed to produce 1 tonne of microalgae and one tonne of oxygen. Assuming 50\% oil content in the dry algae the 1 tonne algae can produce roughly 3.5 barrels of biodiesel \hyperref[b15]{(Kanes, 2009)}. CO 2 produced in the cement industry or in coal fired power plants can be used to fertilize the microalgae production \hyperref[b10]{(Chaput et al., 2012)}. The consumption/fixation of CO 2 takes place during daylight when exposed to the sun, hence it depends on light exposure. It has been reported that over a seven day growth period the microalgae removed 82\% of CO 2 on sunny days (such as Qatar sunny weather), and 50\% on rainy days. Microalgae can consume nitrogen sources 24 h/day. Testing over a seven day period showed that microalgae removed 86\% of NOx with or without light. Microalgae as a feedstock for biodiesel will not compete with food crops because they can be grown on nonarable land (such as Qatar desert).  ear 2012 Y\par
The goal of this research is to investigate the feasibility of producing microalgae oil in Qatar to be used as biodiesel feedstock. To accomplish this goal, the specific objectives were to:\par
The design of the photobioreactor (PBR) is very important in scaling up the growth of microalgae in Qatar's hot sunny desert climate. Open ponds are inexpensive PBRs and easily maintained, but may suffer from contamination. This is less likely in Qatar due to the high salinity of the Gulf seawater. Open ponds have to be shallow to allow for sunlight penetration. Hence a large surface area is needed, which makes harvesting more difficult  {\ref (Scott, 2010)}. Fish tanks with sides blacked out simulate a shallow open pond with a large surface area.\par
Microalgae were grown outdoors in Doha, Qatar during the summer of 2011. This climate provides constant daylight, averaging 95,000 lux (139 W/m 2 ) (Hoki, 1999) whereas the average outdoor daylight in the USA is 50,000 lux or 73 W/m 2 . Qatar is an excellent candidate for growing microalgae because of its high light intensity, desert land, location and CO 2 emissions. Qatar's sunlight intensity will enhance algae photosynthesis. Only 6\% of Qatar is agricultural land, so there is plenty of desert land that could be used to grow algae. Qatar is located in the Gulf, which provides easy access to salt water and is a source of microalgae. Qatar has the highest per capita CO 2 emissions in the world (55.4 tonnes vs. 20 in the US in 2007) due to the petroleum and cement industries. Algae growth is an excellent method of CCU, so Qatar is an ideal candidate for algae growth.\par
The photosynthetic solar constant, which is the yearly mean solar irradiance on the surface of the earth oriented towards the sun above the atmosphere, is 1340 W/m 2 . The photosynthetic active region range is 40-750 nm \hyperref[b5]{(Agrawal, 2010)}. This provides 26\% of the standard solar constant or 350 W/m 2 of input power to the photosynthetic process. The theoretical maximum photosynthetic efficiency is 20\% \hyperref[b9]{(Bonner, 1962)}. Thus, the photosynthetic solar constant, or the maximum output power that can be achieved under ideal conditions would be 70 W/m 2 . The solar biomass constant or theoretical upper limit on the equivalent biomass harvested would be 4.5 mg/m 2 -s.\par
There are three main factors that affect the production cost of algae: (1) cost of land used for the algae production facility, (2) the value of the by-products of algae growth and oil extraction, and (3) the algae oil production rate (g oil/L-day) and hence the importance of this investigation. For open pond algae production systems the capital costs estimates range is \$50,000 to \$250,000 per hectare, and the operating costs are about \$15,000 to \$20,000 per hectare per year. The almost free desert land in Qatar will lower the capital costs estimates. The results of this investigation of using Qatar's hot sunny climate to grow and dry the algae should lower the annual production costs.\par
? Study algae oil production in an open pond system in Qatar's hot sunny desert climate.\par
? Identify if nutrients should be added to the Gulf (off Qatar) seawater to favor algae oil production.\par
? Compare different algae strains to determine the most favorable for algae oil biodiesel feedstock production.\par
Microalgae growth in batch cultures occurs in the following phases: lag, exponential, stationary and death. Ideally, the microalgae should be harvested in the stationary phase because they have reached maximum growth, and the nutrients will have been used up so the microalgae start to accumulate lipids. There are four classes of microalgae; diatoms, green algae, blue-green algae and golden algae \hyperref[b15]{(Kanes, 2009)}. Green algae, e.g., Dunaliella, were used in the present study because they can grow in fresh water and in salt water (such as Gulf seawater). Also, Dunaliella microalgae are the most halotolerant species known and are more tolerant of fuel oil contamination than other species \hyperref[b23]{(Tafreshi. 2009}). Three specific strains of Dunaliella microalgae were used in the present work: Salina, Bardawil and Parva. These strains were chosen for their ability to thrive under hot temperatures in saline water, and their high lipid content (Table \hyperref[tab_0]{2}) ((Abd El-Baky (2004), \hyperref[b20]{Peeler (1989)}, \hyperref[b23]{Tafreshi (2009)}, Scott (2010),  {\ref Fried (1982)}, Ben-Amotz (1990),  {\ref Evans (1982)}). The selected strains were all grown in an open pond system in Doha, Qatar to select the species that exhibits growth, biomass production and oil content.  The experimental work, summarized in Table  {\ref 3}, was done in two phases. The first was to provide enough algae for outdoor testing and establish baseline data on the performance of microalgae in Gulf seawater. These experiments were done indoors using fluorescent lights (4 bulbs illuminating seven 500 mL PBRs) and artificial (contaminants-free) salt water. The second phase aimed at more realistic tests of growing the microalgae in Gulf seawater exposed to Qatar's sunlight and hot climate. This was done outdoors on the roof of a building. The algae growth period for each run was about 14 days. All runs were done in batch mode, i.e., the medium (Gulf seawater with or without nutrients) was placed in the PBR, algae inoculum was added and the air flow was started. Air use had the dual purpose of supplying the CO 2 required for algae growth and oil formation, and providing algae and nutrient medium mixing. Once the algae growth reached the stationary phase they were harvested and dried, then the algae oil was extracted.III. APPROACH \& METRICS a) Experiment Layout Table 3 : Experimental Steps b) 
\section[{Analytical Procedures and Metrics}]{Analytical Procedures and Metrics}\par
Nitrate and pH measurements are important to maintain a consistent growth environment. This was done by collecting 5 mL of algae solution daily and using Mardel test strips. Algae growth was monitored daily by measuring the algae solution absorptivity at 680 nm. This was done by placing the same 5 mL solution in a DR2800 Spectrophotometer. ear 2012 Y\par
The algae inoculum was grown indoors in distilled water using 1.5 M NaCl, macro and micronutrients-this growth media is called "artificial seawater". The measured light intensity of 86,400 lux= 362.9 W/m 2 = photosynthetic photon flux density (PPFD) or light intensity of 1451.5 ?mol/m 2 s ?1 . The carbon source was an air feed from a fish tank air pump.\par
Algae were grown in either just Gulf seawater, or Gulf seawater with added nutrients. The purpose was to determine how the algae would grow in the Gulf seawater, if nutrients are needed, and the species that grew best. Trials without microalgae were included to measure the growth rate of other species in the Gulf seawater. Each trial was exposed to a continuous (24 hour/day) air feed and fluorescent lights.\par
iii. Outdoor Algae Growth Algae were grown outdoors in fish tanks with the sides blacked out so only sunlight would enter from the top. The growth medium had the same salinity as the Gulf seawater (40 g salt/L water), same airflow rate and natural sunlight. Nutrients were added in the same proportions as the algae inoculum growth. Two trials were set up: 20 L Dunaliella Bardawil and 20 L Dunaliella Salina (the results from the indoor lab tests indicated that the Dunaliella Parva does not grow well in the Gulf seawater, so it was not grown outdoors).\par
Figure \hyperref[fig_4]{1} shows the growth measurements of the six different tests of Table \hyperref[tab_1]{4}. The standard trials show very little growth, indicating there was not a strong presence of competing species. During the lag phase there was similar growth in all the microalgae species, but when they reached the exponential phase, the c) Data Analysis/Calculated Indicators The performance of the PBRs was established by calculating the indicators explained in Table \hyperref[tab_3]{5}. Once algae inoculum had grown, the in lab salt water trials were set up (Table  {\ref 6}).\par
Table  {\ref 6} : Algae species/ nutrients for different in-lab trials.\par
Algae were also grown using the Gulf seawater with the same nutrients, airflow and natural sunlight. Two trials were set up:10 L Dunaliella Bardawil and10 L Dunaliella Salina.\par
Algae were dewatered by centrifuging the solution at 5000 RPM for 6 minutes. The saltwater was then removed and DI water was added to clean the algae of salt and the algae were centrifuged again (5000 RPM, 6 minutes). During centrifugation, the algae were completely removed from the water, but the salt remained dissolved in the DI water. This cleaned the algae, and the discarded water did not contain any algae. The algae were dried using natural sun heat by placing the dense algae slurry outdoors (2 hours). The dry algae were massed, then mixed with hexane, heated, and the evaporated hexane was condensed and reused. Over a period of time (2 hours) the hexane extracts the oil in the algae. The algae biomass is removed by vacuum filtration.\par
The filtration was repeated at least four times, or until the filter paper no longer retained algae. Oil is recovered by evaporating the hexane using a hot water bath. These efficiencies are lower than the typical range of photosynthetic efficiency of microalgae. A possible explanation is that the only mixing of the algae was from the air supply bubbling through the solution. This prevents the algae at the bottom of the tank from receiving light as the solution becomes more concentrated with algae, and this will decrease the photosynthetic efficiency. It is evident that the Dunaliella Bardawil grown in DI water with nutrients had the highest photosynthetic efficiency (1.11\%) and carbon sequestration efficiency (6.5\%). Table  {\ref 8} compares the present work results and literature values. The results confirm the feasibility of producing microalgae oil biodiesel feedstock in Qatar and that the process may be further improved. ear 2012 Y Dunaliella Bardawil grown with nutrients in Gulf seawater clearly grew the best. The Bardawil without nutrients showed little growth, and both Parva samples had little growth as well. Due to the poor growth of the Parva, they were excluded from further testing. Instead, it was determined that the Bardawil should be grown outdoors in both artificial seawater and Gulf seawater solutions containing nutrients. Table \hyperref[tab_0]{2} indicates that Dunaliella Salina is promising, so though no indoor testing was performed, Salina was grown outdoors using the same conditions as Bardawil. 
\section[{b) Outdoor Algae Growth and Monitoring Results}]{b) Outdoor Algae Growth and Monitoring Results}\par
Figure \hyperref[fig_5]{2} shows the four outdoor growth runs. It is evident that the trials in artificial seawater had a much longer lag phase than the trials in the Gulf seawater. This could be because the Gulf seawater has its own additional nutrients that expedite the growth of the microalgae. The Dunaliella Bardawil appears to exhibit consistently better growth than the Dunaliella Salina, shown by the higher final absorbance. 
\section[{c) Algae Harvesting}]{c) Algae Harvesting}\par
After harvesting, the algae are massed to determine the production rate, the carbon sequestration and photosynthetic efficiencies. Lipids are extracted to get the algae oil content and the oil production rate.\par
The algae production rate was calculated for the four outdoor trials done in artificial seawater or Gulf seawater with nutrients. The results in Figure \hyperref[fig_6]{3} indicate that the Dunaliella Bardawil shows the highest production, both in artificial seawater (using DI water and nutrients) and in the Gulf seawater off the coast of Qatar. The Dunaliella Salina grew well in the artificial seawater, but there was very little algae production in the Gulf seawater. This indicates that among the algae strains investigated, Bardawil has the highest growth rate in the Gulf seawater with nutrients. The algae production of Bardawil in artificial seawater is 0.02 g/Lday. This is equivalent to about 1.67 g/m 2 -day or 0.0192 mg/m 2 -s. This is less than 1\% of Agrawal theoretical upper limit of biomass harvested of 4.5 mg/m 2 -s. The daily algae production rate of 1.67 g/m 2 -day is lower than the 12 g/m 2 -day reported for Dunaliella Salina in Oilgae.com indicating the possibility of further improvement in the algae production.\par
Figure  {\ref 4} shows the algae oil content results. Dunaliella Salina grown in the artificial seawater with nutrients had the highest oil content. The Dunaliella Bardawil grown in both the artificial seawater and the Gulf seawater had similar oil content. This is indicative that the Gulf seawater provides an environment similar to the artificial seawater, and the microalgae are able to produce oil as usual.\par
The oil production rate of the microalgae is the final oil mass per liter of initial solution per day of growth.\par
Figure  {\ref 5} shows that all three trials were within the same range of oil produced. This further confirms that algae oil as a biodiesel feedstock, can be produced by growing Dunaliella Bardawil in the Gulf seawater.\par
The carbon sequestration efficiency and the photosynthetic efficiency of the microalgae were calculated for the outdoor trials of Dunaliella Salina and Dunaliella Bardawil. Table \hyperref[tab_4]{7} shows that the carbon sequestration efficiency of the microalgae was between 3-6.5\%. The photosynthetic efficiency of the microalgae ranged from 0.053-1.11\%.    \begin{figure}[htbp]
\noindent\textbf{}\includegraphics[]{image-2.png}
\caption{\label{fig_3}Microalgae}\end{figure}
 \begin{figure}[htbp]
\noindent\textbf{1}\includegraphics[]{image-3.png}
\caption{\label{fig_4}Figure 1 :}\end{figure}
 \begin{figure}[htbp]
\noindent\textbf{2}\includegraphics[]{image-4.png}
\caption{\label{fig_5}Figure 2 :}\end{figure}
 \begin{figure}[htbp]
\noindent\textbf{3}\includegraphics[]{image-5.png}
\caption{\label{fig_6}Figure 3 :}\end{figure}
 \begin{figure}[htbp]
\noindent\textbf{45}\includegraphics[]{image-6.png}
\caption{\label{fig_7}Figure 4 :Figure 5 :}\end{figure}
 \begin{figure}[htbp]
\noindent\textbf{2} \par 
\begin{longtable}{P{0.21383647798742136\textwidth}P{0.08018867924528303\textwidth}P{0.1977987421383648\textwidth}P{0.1657232704402516\textwidth}P{0.19245283018867926\textwidth}}
Duna-\tabcellsep Temp\tabcellsep Salinity\tabcellsep Lipid(\%\tabcellsep Found in\\
liella\tabcellsep (°C)\tabcellsep (M NaCl)\tabcellsep dry algae\tabcellsep Gulf salt\\
Species\tabcellsep \tabcellsep \tabcellsep mass)\tabcellsep water?\\
Salina\tabcellsep 22-35\tabcellsep 0.9-4.3\tabcellsep 50\tabcellsep Yes\\
\multicolumn{2}{l}{Bardawil 25}\tabcellsep 3.0-4.0\tabcellsep 30\tabcellsep Yes\\
Parva\tabcellsep 32\tabcellsep 1.5-2.0\tabcellsep 21-25\tabcellsep Unknown\end{longtable} \par
  {\small\itshape [Note: c © 2012 Global Journals Inc. (US)]} 
\caption{\label{tab_0}Table 2 :}\end{figure}
 \begin{figure}[htbp]
\noindent\textbf{4} \par 
\begin{longtable}{}
\end{longtable} \par
 
\caption{\label{tab_1}Table 4 :}\end{figure}
 \begin{figure}[htbp]
\noindent\textbf{4} \par 
\begin{longtable}{P{0.38125469571750564\textwidth}P{0.07535687453042825\textwidth}P{0.19861006761833208\textwidth}P{0.19477836213373403\textwidth}}
Step\tabcellsep Medium\tabcellsep Lighting\tabcellsep Explanation\\
\tabcellsep Phase 1\tabcellsep \tabcellsep \\
1-Inoculum Growth for\tabcellsep Artificial sea water\tabcellsep Fluorescent\tabcellsep Grow inoculum\\
future runs.\tabcellsep \tabcellsep 24 h/d\tabcellsep indoors.\\
2-Indoors Algae Growth\tabcellsep Gulf salt water\tabcellsep Fluorescent\tabcellsep without and with\\
\tabcellsep \tabcellsep 24 h/d\tabcellsep nutrients.\\
3-Indoors non-algae\tabcellsep Gulf salt water -\tabcellsep Fluorescent\tabcellsep Monitor Gulf sea\\
Growth (with and without\tabcellsep no algae\tabcellsep 24 h/d\tabcellsep water without algae\\
nutrients)\tabcellsep \tabcellsep \tabcellsep to check for\\
\tabcellsep \tabcellsep \tabcellsep competing species.\\
\tabcellsep Phase 2\tabcellsep \tabcellsep \\
4-Growth outdoors in a\tabcellsep Artificial sea water\tabcellsep Sunlight (9\tabcellsep Measure solution\\
fish tank PBR with added\tabcellsep and Gulf salt\tabcellsep hours/day)\tabcellsep absorbance daily.\\
nutrients.\tabcellsep water\tabcellsep \tabcellsep \\
5-Algae harvesting\tabcellsep \tabcellsep \tabcellsep Stationary phase\\
6-Algae dewatering and\tabcellsep \tabcellsep Sunlight (9\tabcellsep Centrifuge algae to a\\
outdoor drying using\tabcellsep \tabcellsep hours/day)\tabcellsep slurry. Dry outdoors\\
sunlight\tabcellsep \tabcellsep \tabcellsep to get dry algae.\\
7-Hexane extract oil\tabcellsep \tabcellsep \tabcellsep Solvent extract the\\
then evaporate hexane\tabcellsep \tabcellsep \tabcellsep lipids/oil from algae\\
\tabcellsep \tabcellsep \tabcellsep produced.\\
\multicolumn{2}{l}{Measurement}\tabcellsep \multicolumn{2}{l}{Purpose/ Metrics}\\
\multicolumn{2}{l}{Algae solution absorbance, DR2800}\tabcellsep \multicolumn{2}{l}{Algae growth, from}\\
\multicolumn{2}{l}{Spectrophotometer}\tabcellsep \multicolumn{2}{l}{absorbance/ turbidity}\\
\multicolumn{2}{l}{Acidity of solution (pH),}\tabcellsep \multicolumn{2}{l}{Nutrient content, depletion or}\\
\multicolumn{2}{l}{nitrite and nitrate levels with Mardel test strips.}\tabcellsep \tabcellsep starvation\\
\multicolumn{2}{l}{Algae mass after harvesting and drying (using}\tabcellsep \multicolumn{2}{l}{Algae production rate}\\
\multicolumn{2}{l}{a balance)}\tabcellsep \multicolumn{2}{l}{(g dry algae/L-day)}\\
\multicolumn{2}{l}{Mass of algae oil after extraction}\tabcellsep \multicolumn{2}{l}{Algae oil yield (g oil/100 g dry}\\
\tabcellsep \tabcellsep \tabcellsep algae)\\
\multicolumn{2}{l}{Air flow rate, liters/min (using a rotameter)}\tabcellsep \multicolumn{2}{l}{Carbon sequestration/capture}\\
\tabcellsep \tabcellsep \tabcellsep efficiency\\
\multicolumn{2}{l}{Incident light intensity with a light meter, Lux.}\tabcellsep \multicolumn{2}{l}{Photosynthetic efficiency}\end{longtable} \par
  {\small\itshape [Note: c © 2012 Global Journals Inc. (US)]} 
\caption{\label{tab_2}Table 4}\end{figure}
 \begin{figure}[htbp]
\noindent\textbf{5} \par 
\begin{longtable}{P{0.3555833333333333\textwidth}P{0.49441666666666667\textwidth}}
Parameter\tabcellsep Definition/How Calculated\\
Photosynthetic Efficiency or Light\tabcellsep Ratio of the energy produced by the\\
Capture Efficiency is the light\tabcellsep combustion of the algae to the incident light\\
energy transferred through PBR\tabcellsep ced by the artificial lights used. energy produ\\
and converted to biomass\tabcellsep \\
Carbon Sequestration/Capture\tabcellsep C in dry algae formed/ Total C from the air\\
Efficiency is the mass of C\tabcellsep into the PBR during growth, assuming:\\
sequestered by the algae relative\tabcellsep -Constant air flow rate\\
to the C supplied to the\tabcellsep -Air CO2 (394 ppmv) is the only C source\\
microalgae.\tabcellsep for algae.\\
\tabcellsep -Dry algae are 60.4\% C, based on\\
\tabcellsep literature.\end{longtable} \par
 
\caption{\label{tab_3}Table 5 :}\end{figure}
 \begin{figure}[htbp]
\noindent\textbf{7} \par 
\begin{longtable}{P{0.5280040733197555\textwidth}P{0.003462321792260692\textwidth}P{0.09694501018329939\textwidth}P{0.07963340122199593\textwidth}P{0.12291242362525458\textwidth}P{0.019042769857433806\textwidth}}
\tabcellsep \tabcellsep \multicolumn{2}{l}{efficiencies}\tabcellsep \\
Dunaliella\tabcellsep \tabcellsep Carbon\tabcellsep Photo-\tabcellsep \multicolumn{2}{l}{Photosynthetic}\\
\multicolumn{2}{l}{Algae grown}\tabcellsep Seques-\tabcellsep synth-\tabcellsep \multicolumn{2}{l}{Efficiency relative}\\
outdoors\tabcellsep \tabcellsep tration\tabcellsep etic\tabcellsep to\tabcellsep the\\
\multicolumn{2}{l}{with Nutrient}\tabcellsep Efficien-\tabcellsep Efficien-\tabcellsep \multicolumn{2}{l}{photosynthetic}\\
\tabcellsep \tabcellsep cy* (\%)\tabcellsep cy (\%)\tabcellsep solar\tabcellsep constant\\
\tabcellsep \tabcellsep \tabcellsep **\tabcellsep (\%) ***\\
Salina-\tabcellsep DI\tabcellsep 6\tabcellsep 0.6\tabcellsep 1.1\\
water\tabcellsep \tabcellsep \tabcellsep \tabcellsep \\
\multicolumn{2}{l}{Bardawil-DI}\tabcellsep 6.5\tabcellsep 1.11\tabcellsep 2.1\\
water\tabcellsep \tabcellsep \tabcellsep \tabcellsep \\
\multicolumn{2}{l}{Bardawil-Gulf}\tabcellsep 3.1\tabcellsep 0.053\tabcellsep 0.1\\
saltwater\tabcellsep \tabcellsep \tabcellsep \tabcellsep \\
\multicolumn{6}{l}{*Based on 60.3\% carbon by mass in the microalgae,}\\
\multicolumn{3}{l}{and 394 ppmv CO2 in the air}\tabcellsep \tabcellsep \\
\multicolumn{6}{l}{**Based on microalgae heating value of 5000 cal/g and}\\
\multicolumn{5}{l}{incident light of 89,000 lux, or 130 W/m 2}\\
\multicolumn{5}{l}{*** Based on Agrawal constant of 70 W/m 2}\end{longtable} \par
 
\caption{\label{tab_4}Table 7 :}\end{figure}
 		 		\backmatter   			 
\subsection[{ACKNOWLEDGEMENTS}]{ACKNOWLEDGEMENTS}\par
The authors would like to acknowledge the support of the US-Egypt Science and Technology funds, USDA Contract 58-3148-8-174 and the International Research Opportunity Program (IROP) of the UNH Hamel Center for Undergraduate Research. 			  			 \par
VI. 
\subsection[{CONCLUSIONS}]{CONCLUSIONS}\par
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