A Green Technology for the Production of Biofuels

Dan Whalen

Abstract

. The past several decades have been demarcated by growing concerns about

climate change, a global condition attributed to the emission of green house gases

(GHG). The mitigation of this global threat has been met by the development of green

technologies and alternative fuel sources in an effort to diminish GHG production. One

of the most promising technologies currently being developed is the conversion of

agricultural biomass into alternative fuel sources. Indeed, the conversion of cellulose into

alternative energy sources has become a cornerstone of green technologies. However, the

development of efficient methods for the use of hemicellulose, a major component of

woody materials, in the production of alternative fuels remains elusive. In the present

study, we developed a microbial system capable of converting xylose, the dominant

structural component of hemicellulose, into value­added products. A microbial

consortium grown under various nutrient conditions allowed the efficient metabolic

conversion of xylose into hexoses. Microbes cultured in a standard mineral medium were

found to have the highest conversion rates of xylose to glucose. In contrast, microbes

exposed to nutrient stress preferentially generated mannose. The proficient metabolism

of xylose was attributed to an NADP­dependent xylose dehydrogenase. Intriguingly,

NADP­dependent xylose dehydrogenase displayed a lower activity profile in the nutrient

stress. Thus, we have developed a microbial system capable of converting xylose into

fermentable hexoses. The optimization of this process will enable the production of fuels

in a sustainable manner.

Acknowledgements

I would like to thank Dr Vasu Appanna for his guidance and support throughout

the duration of my project. His willingness to address questions and concerns allowed

me to be confident in my analysis and approach to the project.

My appreciation goes out to Dr. Ryan Mailloux for his patience in guiding me

through a vast array of new procedures. Despite my constant harassment, he continued to

answer my questions with his full attention. Without him , this project would not have

been possible.

Joe Lemire for helping me refine some of my new found lab techniques as well as

providing general support.

My lab mates Steph Leclair, Rami Darwich and Catehrine Kossar for making this

experience as enjoyable and smooth rolling as it was.

My girlfriend, Cara Blasutti, and my parents, Oscar and Kate Whalen for support

through this year which was personally very trying. Without them I would have had to

abandon the project before it started.

Table of Contents ` Page

1.Introduction 5 Global Warming 5 Alternative Fuels 5 Carbohydrates in Lignocellulosic Biomass 6 General Bacteria Metabolism 8 Xylose Metabolism in Soil Bacteria 11

2. Materials 13 2.1 High Phosphate Media 13 2.2 Peroxide Stress 13 2.3 Zinc Stress 14 2.4 Low Phosphate Stress 14 2.5 Bacterial Storage 14 2.6Growing Cultures 14 2.7 Cell Storage Buffers 15 2.8 HPLC Mobile Phase 15

3. Methods 16 3.1 Bradford Biomass Assay 16 3.2 Dubois Carbohydrate Assay 16 3.3 HPLC Analysis 16 3.4 Isolation of Cellular Fractions 17 3.5 Blue Native Polyarylamide­Gel Electrophoresis 17 3.6 In­Gel Staining for XDH Characterization 19 3.7 Bial’s Test for Pentose Concentration 20

4. Results 20 4.1 Bacterial Growth Curve 21 4.2 HPLC Analysis 21 4.3 Carbohydrate Content 23 4.4 Dubois Assay 23 4.5 Bials Test 23 4.6 Gas Chromatography 24 4.7XDH Activity Staining 25 4.8 Zinc and Low Phosphate Cultures 26 4.9 Pentose Concentration in Zinc and Low Phosphate Cultures 27

5. Discussion 27 5.1 Biomass 28 5.2 Spent Fluid Analysis 28 5.3 Dubois Assay 29 5.4 Pentose Concentration 29 5.5 Gas Chromatography 30 5.6 Xylose Dehydrogenase Characterization 31

6. Optimization and Further Research 34

List of Figures Page

Figure 1: Glycolysis………………………………………………………………….9

Figure 2: The Citric Acid Cycle……………………………………………………..10

Figure 3 : Proposed Pathway for D­Xylose Metabolism……………………………12

Figure 4 : Bacterial Growth Curve in 20mM Xylose………………………………...21

Figure 5: Chromatogram of Spent Fluid HPLC……………………………………...22

Figure 6: Results of Dubois Assay…………………………………………………...23

Figure 7 : Bial’s Test Results………………………………………………………….24

Figure 8: XDH Activity Stains………………………………………………………..26

Figure 9: Bacterial Growth Curve for Zinc and Low Phosphate Cultures……………27

Figure 10: Pentose Concentration in Zinc and Low Phosphate Cultures……………..28

Figure 11: Metabolic Pathway for D­Xylose Metabolism In Soil Bacteria…………...29

List of Tables Page

Table 1: Conversion Rates of Xylose into Hexosans…………………………………25

1. Introduction:

Global warming has become one of the most severe environmental issues in recent

history. The impact of climate change will be severe if measures are not taken to reverse

and or slow the mechanisms contributing to the global increase in temperature.

Greenhouse gas production is the greatest contributor to the rise in global temperature.

The accumulation of greenhouse gases (GHG) in our atmosphere has caused the heat

waves produced as the suns radiation strikes the earth to become trapped inside the

earth’s atmosphere (Crowley 2000). CO2 emissions produced from hydrocarbon

combustion in automobiles, coal power stations, and aircraft account for the damaging

effects of GHG, Thus, cleaner fuels and alternative energy sources are required to

diminish GHG levels. While nuclear and wind power generators are replacing the carbon

dioxide producing coal power stations, transportation is in need of a major overhaul in

order to meet the environmental needs of the future.

Alternative Fuels

Current projects underway which are capable of replacing hydrocarbon burning

vehicles include hydrogen fuel cells, as well as alcohol fueled engines. Hydrogen is

possibly the cleanest initiative proposed as it uses the energy stored in hydrogen to power

the vehicle with H20 as the major waste product. However, issues such as safe

transportation due to hydrogen’s highly combustible nature as well as sufficient supply

are hurdles in its acceptance into the mainstream infrastructure (Ashley 2005). Ethanol

on the other hand is perhaps the leading liquid fuel replacement for hydrocarbons as it

produces very little carbon dioxide as a bi­product. The benefits of switching to ethanol

based fuel supplies are that very little needs to be done to either current automotives or

the infrastructure which supplies them to make the switch to ethanol ultimately happen.

This is because ethanol is both less volatile than hydrogen and can increase the efficiency

of hydrocarbon combustion (Lee 1997). The major argument against ethanol

development as a major fuel source is that there exists insufficient supply resources for its

production. Indeed, current methods for ethanol production involve using the starch from

corn as the glucose source for ethanol fermentation. The problem with this process is

that the yield of corn required for mass scale production is simply far too high. Corn has

a relatively low energy per mass ratio meaning that in order to supply the world’s

growing energy needs; corn would have to be grown over the entire earth’s available

surface (Sun and Cheng 2002). However, alternative sources of fermentable sugars are

now being employed to fill the energy gap. Other fermentable sugar sources include

cellulose and hemicellulose. Butanol has also become an interesting prospect due to its

higher energy density than ethanol and lower hydroscopicity. Butanol can be produced

using metabolic and genetic engineering techniques on bacteria such as Escherisia coli in

order increase the formation of Butanol (Atsumi et al 2008). Ethanol has been criticized

somewhat because a method for storage which nullifies its higher hydroscopicity has not

yet been found. Some techniques have been devised as of late which are approaching

viability, involving creating butanol and other high carbon alcohols using Escherechia

coli along with some non native enzymes (Atsumi et al 2008).

Carbohydrates from Lignocellulosic Biomass

Cellulose is the most abundant carbohydrate on the planet and is therefore an

attractive prospect for the production of biofuels. Cellulose is simply a polymer of D­

glucose which can be harnessed to generate ethanol efficiently using glucose fermenting

bacteria (Lin and Tanaka 2005). Cellulose is the major structural component of plant cell

walls, wood, and bark making this D­glucose polymer the greatest renewable energy

resource on the planet. This makes biomass resources easily available in many rural

areas, and consequently may also help play an important role in providing jobs in rural

communities if it can be efficiently utilized as a fuel source (Lin and Tanaka 2005). The

first step in the preparation of lignocellulosic biomass for ethanol production is to

delignate it to free the polymers of cellulose and hemicellulose from the binding agent

lignin. Treatment of wood with chlorous acid yields a holocellulose mixture which upon

treatment with potassium hydroxide will produce a soluble hemicellulose/cellulose

mixture. The cellulose must then be broken apart with either chemical hydrolysis or

biological means using the enzyme cellulase (Hahn­Hagerdahl et al. 1994). The

liberated D­glucose can then be fermented using Sachromyces cerevisae or other

fermentative bacteria to produce ethanol (Sedlack and Ho 2004). The fermentation of D­

glucose has been performed efficiently for many hundreds of years as this fermentation

process is nearly identical to that used by beer and wine makers. The ethanol can then be

further distilled in order to be made properly combustible. Lignocellulosic biomass also

contains hemicellulose, a complex polymer of hexosans and pentosans. Hemicellulose is

closely associated with lignin forming a major structural moiety in higher plant tissues.

The structure of Hemicellulose is composed of a D­xylose backbone with side chains of

arabinose, hexoses and modified carbs. Roughly 30% of the dry mass of lignocellulosic

biomass is actually the pentose sugar containing polymer Hemicellulose. 30% of

hemicelluloses sugars in hardwood trees is D­Xylose, a number which can reach 70% in

many types of straw, sugarcane bagasse and soybean stalks (Lin and Tanaka 2006).

Current methods are being studied using xylose fermenting bacteria to maximize ethanol

production and efficiency. The presence of D­xylose along with D­glucose in the

hydrolyzed product presents biochemical problem when trying to directly ferment both

simultaneously. This is because the glucose inhibits the xylose fermentation and the

xylose inhibits the fermentation of glucose when using recombinant fermentative bacteria

(Sedlack and Ho 2004). Indeed, the inability o fermenting microbes to properly ferment

xylose and other pentosans makes the use of agricultural biomass for biofuel production

inefficient. Thus, it is important to find more economically viable methods of converting

xylose into value added products in order to increase the overall efficiency of

lignocellulotic biomass fermentation.

General Bacterial Metabolism

Bacteria are the key to finding cheap efficient ways to metabolize wood

carbohydrates because evolution has allowed for the broad development of their

metabolic pathways over millions of years. Bacteria rely on various metabolic pathways

in order to convert various carbon sources into building blocks and ATP. Carbohydrate

metabolism in bacteria involves a complex network of metabolic reactions which include

parts of glycolysis, gluconeogenesis, the citric acid cycle, as well as the Pentose

Phosphate Pathway and other intermediate shunts. Glycolysis is the initial phase in the

production of energy from glucose and involves a series of 10 enzyme catalyzed steps

which conclude with the production of the 3 carbon pyruvate molecules. The first stage

of glycolysis involves the conversion of D­ glucose into the trioses, glyceraldehyde­3­

phosphate and DHAP. This conversion requires the phosphorilation, isomerization, and

aldol cleavage of D­glucose. Stage 2 of glycolysis involves the NAD­dependent

oxidation of the trioses, a process coupled to the generation of 4 ATP. In order to

maintain glycolytic flux the NADH is oxidized by the conversion of pyruvate to ethanol,

a process called alcoholic fermentation. Alcoholic fermentation is required to maintain

glycolytic efficiency when O2 is scarce. However, when O2 is available, NADH is

oxidized by the electron transport chain. (Horton et al).

Fig 1 Glycolysis

Gluconeogenesis is basically the reverse of glycolysis. When ATP production is

no longer required pyruvate is used to fuel the production of nucleotide and carbohydrate

precursors needed for biosynthetic reactions. The 3 enzymes which are not homologous

between both glycolysis and gluconeogenesis are the enzymes which modulate the rate of

the reactions in either direction and are controlled indirectly by the ATP levels inside the

cell. If ATP reserves in the cell are lower than required however, the pyruvate produced

during glycolysis is brought into the mitocondria where it is decarboxylated by pyruvate

dehydrogenase to form Acetyl CoA. The Citric acid cycle then combines a molecule of

oxaloacetate which is formed as a product of the previous cycle to form citrate. The citric

acid is then systematically oxidized by the concerted action of TCA cycle enzymes

generating NADH, FADH2, and evolving CO2.

Fig 2: The Citric Acid Cycle

The NADH and FADH2 donate their electrons to the respiratory complexes. The

electrons are then passed down their electrochemical gradient to the terminal electron

acceptor O2. This generates the H+ gradient required to drive ATP production. Bacteria

can also produce acetyl CoA from lipids by cleaving the fatty acids from the glycerol in

stored lipids with phosphodiesterases to produce 3 fatty acid chains which are essentially

chains of Acetyl groups which are easily converted to acetyl CoA(Horton et al 2006).

These two pathways are key in the production of energy in all non photorespiring cells,

both eukaryotic and prokaryotic.

Xylose Metabolism in Soil Microbes

Soil microbes have been shown to exhibit some variation in the metabolism of

both xylose and arabinose. Since Xylose is a major constituent of agricultural biomass,

the aim of this project is to develop a microbial system capable of converting D­xylose

into value added products. If the bacteria are metabolizing bacteria for energy

production, xylose tends to follow either the proposed Weinberg or Dahms pathway. The

intial step in both the Dahms and the Weinberg proposed pathways is the oxidation of

xylose into xylonolactone by xylose dehydrogenase, while reducing one molecule of

NAD+ in the process. In the next step D­ Xylonate is produced through the use of the

enzyme xylonolactase. Subsequent removal of a molecule catalyzed by Xylonate

Dehydratase yields 2 keto 3 deoxyxylonate and it is here where the two proposed

pathways begin to differ (Stephens et al 2006). In the weinbeg pathway the xylonite is

dehydrated and then oxidized before becoming aKG which enters the TCA cycle for

either energy production or so it can exit as malate to be further transformed for storage

purposes as in fig 1. The Dahms pathway involves utilizing the 2 keto 3 deoxypentose as

a substrate to produce one molecule of pyruvate and one glycoaldehyde molecule.

Fig 3 Proposed pathway fro D­Xylose Metabolism

If the bacteria cell determines that ATP levels inside the cell are adequate it can

then shift its metabolism to produce carbohydrates such as glucose and mannose in a

more direct manner by utilizing reactions of the Pentose phosphate pathway (PPP). This

is the common mechanism utilized by yeasts to metabolize xylose into ethanol. In this

proposed mechanism the xylose is transformed into xylitol by using nadp+ dependent

xylsoe reductase. Xylitol is then converted to xylulose 5 phosphate, a substrate for PPP

degredation through subsequent oxidation and phosphorylation reactions . The xylose 5

phosphate then enters the Pentose phosphate pathway to yield pyruvate which can either

be used for anaerobic energy production yielding ethanol, or sequestered for glucose

production through the gluconeogenic pathway. The conversion of xylose to glucose

would also be an economically viable process as glucose can easily be fermented into

ethanol as well. By producing additional ethanol or glucose from the hemicellulosic

components of lignocellulose the overall viability of utilizing biomass would increase

considerably.

The media in which any organism is grown accounts for the specificity of the

metabolic pathways utilized by that organism both for energy production and energy

storage purposes. By determining experimentally the preference of specific bacteria for

producing a desired product it is possible to maximize the production of that product.

The reason for this is that the buildup of metabolites, reactive oxygen species, and other

chemicals can change the ability of various enzymes in the cell to deal with their

substrate. In this experiment I will be growing cultures of soil bacteria in a variety of

control and nutrient stress environments in order to analyze and maximize the production

of value added products such as keto acids, pyruvate, butanol, ethanol and glucose. I

will be monitoring the growth rate and protein concentration at confluency of the bacteria

in each media using Spectrofluorometric techniques. The metabolic components of the

media at various timepoints will also be analyzed using various methods. Some enzyme

analysis will also be performed in order to try and determine the metabolic direction of

both control and stress media cultures in an effort to steer the metabolism of the organism

towards whichever value added metabolite is being produced. My goal is to produce in

an efficient manner a metabolite from xylose which could offset the cost of ethanol or

butanol production, both considered to be viable alternatives for the dwindling oil

reserves and increased GHGs of our world.

2. Materials

Microbial growth conditions and media

High phosphate control media was prepared by adding 12.0g Na2HPO4, 6.0 g KH2PO4,

0.4 g MgSO4∙7H2O, 1.6 g NH4Cl, and 1 mL of a trace element solution to 600 mL of

ddH2O. Trace element solution contained 2 μM FeCl3∙6H2O, 1 μM MgCl2∙4H2O, 0.05

μM Zn(NO3)2∙6H2O, 1 μM CaCl2, 0.25 μM CoSO4∙7H2O, 0.1 μM CuCl2∙2H2O, 0.1 μM

NaMoO4∙2H2O. The pH of the trace element solution was adjusted to 2.75 with dilute

HCl, and the solution was stored at 4 o C. The pH of the trace element solution was

adjusted to 6.8 with dilute NaOH and the volume was brought to 360 mL with ddH2O.

After stirring and bringing the pH of the phosphate media to 6.8 with 2M HCL, the final

volume was brought to 2L by adding ddH20. The media was subsequently autoclaved at

121 o C for 60 min and allowed to cool to room temperature. Xylose Stock was prepared

by adding 6.0g of D­Xylose to 200ml of ddH20 and then stirred. The Xylose stock was

sterilized using a disposable vacuum unit with a pore size of 0.2 microns to sterilize.

20ml of 20mm Xylose and 180 mL of High Phosphate media were then added to a

500mL Erlenmeyer flask. The final concentration of Xylose in the control growth media

was 20mm.

Peroxide Stress

The peroxide media was prepared as above however 200ul of 100mM peroxide was

added to the 500ml Erlenmeyer flask along with 180ml of High Phosphate media and

20ml of Xylose stock.

Zinc Stress

Zinc media was prepared by adding 400ul of 10mM Zinc to 180ml of High Phosphate

Media and 20ml of Xylose Stock.

Low Phosphate Stress

Low phosphate media was prepared by adding100x less Na2HPO4. The media therefore

included 0.12g Na2HPO4, 6.0 g KH2PO4, 0.4 g MgSO4∙7H2O, 1.6 g NH4Cl, and 1 mL of

a trace element solution. The volume of the media was adjusted to 2L and sterilized as

above. The Xylose Phosphate media contained 180ml of low Phosphate media and 20ml

of Xylose stock.

Bacterial Storage

Xylose slants were prepared by 270ml of High Phosphate Media and 4.8g of Agar to a

500ml beaker and was stirred over heat until it dissolved. The solution was then

autoclaved for 60 min at 121 o C in order to sterilize. 30ml of Xylose stock was then

added to the warm solution and 10ml portions of the agar solution was poured into sterile

test tubes. The slants were allowed to harden with the test tubes lying at an acute angle

inorder to maximize the surface area of the hardened gel. After the slants were cooled,

they were then refrigerated at 4 degrees Celcius. Slants were then inoculated every time

a new culture was started and allowed to grow for 4 days until visible growth was

observed. The inoculated slants were then refrigerated at 4 degrees Celcius.

Growing Cultures

Precultures were made adding 90ml of Phosphate media and 10ml of Xylose stock into a

250ml Erlenmeyer flask. The media was then inoculated with a streak from a fresh

Xylose slant incubated for 48 h in a water bath shaker (model G76, New Brunswick

Scientific) at 26 o C and 140rpm. 1mL of pre­culture was then isolated and added to

200ml of each of the 4 different stress media in a 500mL Erlenmeyer flask. These

cultures were again allowed to grow in the water bath shaker and 10ml samples were

isolated at 8h post inoculation and then 24h intervals up to 96h and refrigerated at 4

degrees Celsius. Biomass isolation was performed by subsequent centrifugation of the

samples at 4000 rpm for 30 min and removal of the supernatant. The pellet was then

resuspended in 1ml of NaOH.

Cell Storage Buffer (CSB) with DTT

Cell storage buffer was made by adding0.03084 g Dithiothreitol (1 mM), 1.211 g

Trizma base (50 mM), 0.035 g Phenylmethylsulfonylfluoride (1 mM). DTT and Trizma

base to 80 mL of ddH2O and heated until fully dissolved. PMSF was then added and the

solution was cooled and brought to pH = 7.4 with 2 M HCl. Total volume of 200 mL was

achieved by adding of ddH20. The buffer was then covered and refrigerated at 4 º C.

Tris Reaction Buffer

Reaction buffer was made using (5 mM) MgCl2 and 25 mM Tris in a final volume of 1 L.

Tris and MgCl2 were added to 500 mL ddH2O. A pH of 7.4 was then achieved using 2M

HCl. Final volume was then brought to 1 L using ddH2O.

HPLC Mobile Phase

HPLC mobile phase was prepared by adding 2.72 g of 20mM (≥99.9%) KH2PO4 to 500

mL ddH2O. Mobile phase pH was adjusted to 2.9 with 2 M HCl.

3. Methods

Determination of Biomass

Biomass was monitored using the Bradford Assay. 10mL isolated samples were

centrifuged at 4000rpm for 30min. The supernatant was removed and saved for HPLC

analysis. The pellets were resuspended in 1ml of NaOH, placed in microcetrifuge tubes,

and boiled for 5 minutes in order to degrade the cell membranes and denature proteins.

1.5 mL cuvettes were used for the assay. Samples were diluted in the cuvette with 200uL

of Bradford reagent. The volume was then brought to 1mL with ddH2O and allowed to

stand for 5 minutes. The absorbance was then measured at 595nm using a

spectrophotometer. BSA, at a final concentration of 10ug/mL was used as the standard.

Dubois Carbohydrate Assay

Total carbohydrate concentrations were determined using the Dubois

carbohydrate assay. 800 µL of ddH2O, 200 µL of sample supernatant, 1 mL of phenol

(5% w/v) and 5 mL of H2SO4 (90% v/v) were mixed in 10 mL test tubes. Absorbance

was measured at 488 nm by transferring 1 mL of each reaction mixture into 1.5 mL

cuvettes. Standard curves were developed using D­glucose varying from 0.1 to 1mg/mL.

Bials Test for Pentose Concentration

In order to measure the consumption of Xylose and formation of Pentose sugars

in the culture media, a Bial’s Assay was performed. Bial’s reagent was prepared by

adding 0.3g of orcinol and 0.05 g of ferric chloride to 100mL of 12M HCl. A standard

curve was created adding 0.01ml, 0.02 ml, 0.03ml, 0.04ml, and 0.05ml of 1mg/ml xylose

stock to 5 different test tubes. 1ml of Bial’s reagent was then added to each test tube and

boiled for 5 minutes in a fume hood. The contents of each test tube were then transferred

to 1.5 ml cuvettes and their absorbance measured at 660nm to measure the formation

furfurol which forms a blue green product in the presence of orcinol and ferric chloride.

Samples were prepared in triplicate by adding 150ul of the spent fluid of 8, 24, 48, 72, 96

and 120 hour isolations to small test tubes. 3ml of Bial’s reagent was then added to each

test tube and the tubes were heated in a boiling water bath for 5 minutes. The absorbance

was again measured at 660nm. Both control and stress media were tested.

Metabolic analysis and HPLC

The supernatant of all samples were separated using a Phenomex Reverse Phase

C­18 column with an injection volume of 10 µL and a flow rate of 0.7 mL/min. The

separations module is coupled with a UV/VIS spectrophotometer that was calibrated at an

absorbance of 240 nm in order to pick up organic acids. Samples were prepared for

HPLC by running 2mL of spent fluid through 1mm of cotton in a Pasteur pipette .

Samples were then injected into the HPLC column.

Subcellular fractionation and preparation for Blue Native PAGE

Cell cultures were grown for 30h and then poured into centrifuge tubes. After

centrifuging for 30 min at 4000rpm the supernatant fractions were removed and the

pellets were resuspended in 0.85% NaCl and then centrifuged again at 4000rpm for

another 30min. The supernatant fractions were discarded and the pellets were

resuspended in 500ul of CSB. Low intensity sonication was performed (Brunswick

Ultrasonic Processor) with four cycles of 15 s at power level ≤4 on ice. This was done to

lyse the cell membranes and liberate to separate cytoplasmic and membrane bound

proteins. The lysate was then separated into membrane and soluble fractions by

centrifuging at 50 000 rpm in 40º C for 2h. The soluble fraction was decanted and placed

in a micro centrifuge tube. The pellet was resuspended in minimal of CSB. Protein

concentrations of both fractions were determined using the Bradford method. 133ul of 3x

BN buffer, 50 ul of 10% malticide, sufficient fraction to give a final protein concentration

of 125 ul/ml were put into a microcentrifuge tube and the final volume brought up to

500uL with ddH20. Membrane fractions and 500ul of soluble fractions were then frozen

at ­21ºC until needed fro Blue Native Page electrophoresis.

Blue Native Polyacrylamide Gel Electrophoresis (BN­PAGE)

The Bio­Rad MiniProtean 2 electrophoresis system was used for running all of the gels.

Table 2 outlines the contents of all polyacrylamide gels. The separating gel was

composed of 2.9 mL of 4% acrylamide solution and 2.9 ml of 16% acrylamide solution.

4­16% linear gradient gels were generated using a gradient former (BioRad) and a

peristaltic pump. This type of Bio­Rad gradient allows for broad range separation, with

1mm spacers being used in all cases. After all the gel was in the form the gel was

overlaid with isopropanol to allow for proper polymerization. After waiting 20 minutes

for solidification of the separating gel, the isopropanol was absorbed from on top. The

stacking gel ingredients were then mixed together and inserted on top of the stacking gel

with a pasture pipette. While the stacking gel was still liquid, a plastic comb was inserted

to allow for the formation of wells. After polymerization of the stacking gel the combs

were removed and 60ug of protein was applied to the wells and subsequently overlaid

with cathode buffer (see table 3 for BN­Page buffers). The gels were then placed in a

electrophoresis tank with the inner portion of tank being filled with blue cathode buffer

and the outer tank was filled with anode buffer. The tank was then placed in the

refrigerator at 4ºC and a voltage of 80V was applied between ends of the gel until the

protein had migrated into the separating gel. Upon the proteins reaching the separating

gel the voltage was increased to 200V. Once the proteins had migrated halfway through

the separating gel the blue cathode buffer was replaced with colorless cathode buffer.

Once the running front had reached the bottom of the gel the electrophoresis was halted.

Reagent 4% Gel 16% Gel Stacking Gel 49.5% Acrylamide 243 937 273 3x BN Buffer 967 967 1136

ddH2O 1699 223 2000 75% Glycerol 0 773 0 10% APS 9.7 7.6 30 TEMED 1 0.8 2.5

Ingredients for 4%­16% BN­PAGE Gels. All volumes are in µL.

Solutions for BN­PAGE

Blue Cathode Buffer (1L) 8.96 g Tricine (50 mM) 3.138 g BisTris (15 mM) 0.2 g Coomassie blue G 250 pH 7.0 at 4 o C

3x BN Buffer 9.84 g Aminocaproic acid (1.5 M) 1.567 g BisTris (150 mM) pH 7.0 at 4 o C

Coomassie Brilliant Blue Staining Solution 50% Methanol 10% Glacial acetic acid 0.2% Coomassie Blue R 250

Colourless Cathode Buffer (1L) 8.96 g Tricine (50 mM) 3.138 g BisTris (15 mM) pH 7.0 at 4 o C

Anode Buffer 10.45 g BisTris (50 mM) pH 7.0 at 4 o C

Destaining Solution 50% Methanol 10% Glacial acetic acid

In­Gel Stain for Xylose Dehydrogenase Characterization

Activity of Xylose Dehydrogenase (XDH) was visualized by separating

membrane cell fractions on a 4%­16% BN­PAGE gel and incubating the gel in 4 different

reaction mixtures. All reaction mixtures contained 250ul phenozine methyl sulfate

(PMS), 500ul of Iodonitrotetrazolium (INT), and 75ul of 200mM xylose. In order to

characterize the enzyme XDH, I then varied the concentration of two cofactors, NAD+

and NADP+. One mixture was 0.5mM NAD, one was 0.1 mM NAD, one was 0.5mM

NADP and one contained 0.1mM NADP. Gel slabs were incubated in 3mL of reaction

mixture per dual lanes of control and stress. Formazan precipitate forms directly over the

site of enzyme catalysis with the amount of precipitate being proportional to enzyme

activity. Upon visualization, the reaction is quenched by transferring the gel slab into

destaining solution.

4. Results

As shown in figure 4 both the peroxide stressed and control cell cultures

reached confluency in roughly 30 hours. However, the biomass of peroxide stress was

less than control. The biomass measurements were for total protein content in the cell

fraction of the media and were measured using the Bradford method. Upon reaching

confluency, the biomass in both cultures remained relatively stable.

Bacterial Growth Curve in 20mm Xylose Media

­0.05

0

0.05

0.1

0.15

0 24 46 72 96 120

Time Of Isolation(hrs)

Bio

mas

s (T

otal

Prot

ein,

mg/

ml)

Control

Peroxide Stress

Fig 4 :Bacterial Growth in 20mM Xylose Media

HPLC Analysis

Keto Acids are a valuable commodity to the medical community thus we

deciphered if xylose was being converted into either pyruvate or aKG.

Figure 5: Chromatogram of 20mM Xylose Control (left) and Peroxide stress media c

(right) cultures spent fluid

HPLC analysis of the media revealed the presence of pyruvate from the peroxide stressed

cultures. Indeed a small peak at 5 minutes was recorded in the peroxide stressed media.

No peak was observed in the control. However, the spike was not significant; in fact the

absorbance spike of the pyruvate did not even match the peak in absorbance due to the

mobile phase. After calculating the integral of the area underneath the curve at 5 minutes

the concentration of pyruvate in the stress culture was found to be in the microgram per

milliliter range. The results from the HPLC spent fluid analysis demonstrated an overall

lack of buildup with regards to metabolic intermediates.

Measurement of Carbohydrate Levels

pyruvate

0.008

0.001

0.004

2 4 6 8 10

A

Minutes

pyruvate

0.008

0.001

0.004

2 4 6 8 10

A

Minutes 4 10

0.008

A

minutes

0.001

4 10

0.008

A

minutes

0.001

A Dubois assay was performed to measure the total carbohydrate content in the

spent fluid of both control and peroxide stressed cells. Both cultures showed a slight rise

in carbohydrate concentration after the low point at 24hr, followed by a stable period of

little to no change in carbohydrate levels. The control media however showed a sharp

rise in carbohydrates after 120 hours.

Total Carbohydrate Content in Spent Fluid of 20mM Xylose Cultures

0

0.1

0.2

0.3

0.4

24 48 72 96 120

Time of Isolation

Carbo

hydrate

Con

centratio

n (m

g/mL)

control stress

Figure 6: Results from Dubois assay showing relatively stable carbohydrate

concentrations with low points at 24hr isolation. Control media had an increase in

total carbohydrates at the 120 hour mark.

A Bial’s test was then performed in order to detect the levels of pentose sugars in

the mixture. Results show that the low point in pentosans occurs around the 24 hours of

growth point. This is coincides with a time to confluency for both cultures of around 30

hours. After the 24 hour point there was some increase in pentose concentration as

xylose is evidently converted into other pentoses. The concentration of pentose sugars

goes from 3g per Liter at the time of inoculation, all of which is xylose, to a low

concentration of 1g per Liter after 24hours in both culture media. These data indicate

that xylose is consumed quite rapidly since the concentration of xylose at time 0 was

3g/mL. Furthermore, these data suggest that xylose is converted into hexose sugars.

Thus, we setout to identify which hexoses were being produced.

Pentose Concentration

0

0.005

0.01

0.015

0.02

0.025

8 24 48 72 96 120

Time of Isolation (Hours)

Pen

tose Con

centratio

n (m

g/ml)

control stress

Figure 7: Bial’s test results showing a depletion of pentosans after 24 hours

and a slight increase after 120 hours.

Isolations from both cultures were taken after 24 hours and the spent fluid was

sent to Paprican labs for carbohydrate analysis. The lab used gas chromatography to

distinguish between carbohydrates in the mixture. Results indicated that almost all of the

xylose in both control and stress media was consumed after 24 hours. In the control

media the xylose concentration went from 20mM to 1.6mM at the 24 hour isolation and

the concentration after 24 hours in the stress media was 1.7mM. The overall

carbohydrate content in the mixture was still quite high remaining at 10mM. The

conversion of Xylose into other sugars was apparent with glucose concentration of 6mM

in the control culture and 4.5mM in the stress media after 24 hours of incubation. The

remainder of the carbohydrates in the spent fluid was mannose with concentrations of

2.6mM for the control and 2.9mM in the peroxide sample. The level of glucose and

mannose produced are significant given the market value and ethanol yielding properties

of these two carbohydrates. Table 1 provides a summary of xylose to hexose conversion

rates.

Type of hexosan/Culture Xylsoe to Mannose Xylose to Glucose

Control 10% 29%

Stress 14% 18%

Table 1: Conversion rates from xylose into hexose sugars.

Xylose Dehydrogenase Activity Staining

Blue­Native PAGE was performed on membrane fractions form both control and

stress cultures. After separating the protein, activity stains were performed in order to

test fro activity of XDH under different concentrations of either NAD + or NADP+

cofactors. This was performed in order to better characterize xylose dehydrogenase. The

consumption of NAD+ would indicate that xylose dehydrogenase could directly impact

the aerobic respiration of the cell. A strict dependence on NADP+ could play an

important role in the ability of the bacteria to control reactive oxygen species as NADP+

is essential not only in biosynthetic reactions but in anti­oxidant defense.

Fig 8 Xylose dehydrogenase activity stains with 0.1mM nadp+ (top left), 0.5

mM nadp (top right), 0.1mM nad+ bottom left, and 0.5 mM nad+ (bottom right).

Control membrane fraction in left lane, stress membrane fraction in right.

The activity of XDH was greatest in the reaction mixtures containing NADP+ as a

cofactor. The reaction mixture containing 0.5mM NADP+ showed the most substantial

activity bands, however even at concentrations of 0.1mM nadp+ the activity of XDH was

quite evident. The 0.1 mM NAD+ showed very little activity whereas the 0.5mM NAD+

reaction mixture showed some moderate activity. In all reaction mixtures the activity

bands were more evident for the control fractions then for the stress fractions.

Zinc and Low Phosphate Cultures

Zinc and low phosphate cultures were also started and their biomass observed to

determine if these cultures would be viable in the production of value added products

especially monosacharides due to the tendency of these stresses to increase

gluconeogenesis. Results showed a slower time to confluency than the other two cultures

and consequently I focused mainly on the control and peroxide stress cultures from that

point on.

Bacterial Growth in 20mM Xylose

0 0.02 0.04 0.06 0.08 0.1 0.12 0.14 0.16 0.18

24 48 72 96

Time of Isolation (hrs)

Biomass (m

g/ml o

f total

protein)

zinc low phosphate

Fig 9 Bacterial Growth Curve for Zinc and Low Phosphate Stress cultures

Near the end of the project I did however have time to perform a Bial’s test on the

zinc and low phosphate cultures. Results showed a low level of pentose production later

on in the growth phase which may indicate production of hexosans or keto acids. Further

investigation should be performed in order to better analyze the metabolism of the

bacteria in these cultures.

Pentose Concentration in Spent Fluid

0 0.01 0.02 0.03 0.04 0.05 0.06 0.07 0.08

24 48 72 96

Time of Isolation

Pen

tose Con

centratio

n (m

g/ml)

low phosphate zinc

Figure 10 Pentose Concentration in low phosphate and zinc stressed cultures

Discussion

The formation of economically viable products from xylose is an important step

in bringing lignocellosic biomass to the forefront of alternative fuel technology. The aim

of this project was to try to find an efficient method for converting xylose into value

added products such as ethanol, butanol, glucose or keto acids by manipulating the xylose

containing media. 20mM xylose cultures were grown in control, peroxide, zinc and low

phosphate mineral mediums, although much of the focus was placed on the control and

peroxide stress cultures. It was identified that much of the xylose was converted to

hexoses in both control (39 percent) and peroxide (32 percent) cultures. Production of

organic acids in the spent fluid was insignificant.

Biomass

The biomass was measured using the Bradford method. Growth rates for control

and peroxide cultures were relatively similar as both reached confluency in just under 30

hours. Protein concentration for peroxide cultures at confluency was slightly lower than

in the control media with protein concentrations of 1.2 and 1.3mg/ml respectively. After

reaching confluency the biomass in each culture remained quite stable. The oxidative

stress caused by the peroxide in the media was the most likely contributor to the lower

cellular levels at confluency as high levels of peroxide is know to be caustic.

Hydrogen Peroxide was chosen as a stressor due to its ability to increase cellular levels of

ROS. These toxic moieties have been shown to perturb various cellular structures and

functions. For instance, ROS are capable of altering various enzymes within the TCA

cycle and possibly causing the buildup of keto acids in the culture media. Thus,

hydrogen peroxide is an excellent candidate for modifying the molecular pathways of

xylose metabolism.

Spent Fluid Analysis

HPLC was used to analyze the spent fluid of both cultures at 24 hour increments.

The absorbance of the eluate was measured at 210nm in order to detect the presence of

organic acids such as pyruvate and alpha keto glutarate. I hypothesized the buildup of

organic acids in the peroxide stressed media as ROS have been shown to inhibit TCA

cycle enzymes such as akG dehydrogenase. Results from the HPLC analysis showed that

there was however very little buildup of either pyruvate or akG, suggesting that either the

TCA was functioning optimally or the metabolism of the bacteria was shifted mostly

towards gluconeogenesis. In the peroxide stressed cells there was a small peak after 5

minutes, however the peak did not even manage to eclipse that of the mobile phase which

eluted from the stationary phase after 4 minutes. It was clear that in both control and

stress cultures the metabolism of the bacteria was not geared to produce organic acids in

high concentrations.

Dubois Assay

The next step in the metabolic analysis of microbes in the two cultures was to

measure the carbohydrate levels in the spent fluid. The Dubois assay itself measures only

the presence of aldehyde groups which are present in both pentose and hexose sugars.

Both control and stress showed relatively stable carbohydrate concentrations in their

media from inoculation to 120 hours post inoculation time points. Since it was obvious

that xylose was being consumed as the bacteria proliferated, it was clear that there was a

production of another carbohydrate in the growth media. The production of glucose,

mannose or other carbohydrates in the growth media would account for the overall levels

of carbohydrates in the growth media being stagnant. At the 120 hour time point, a spike

in carbohydrate levels in the control media was observed which may have been due to a

gluconeogenic shift in the overall metabolism in the cell causing a buildup of glucose in

the spent fluid.

Pentose Concentrations

In order to better understand the rate at which pentoses were being consumed as

well as produced, a Bial’s test was performed. The production of furfurol from pentose

which produces a blue green color in the presence of iron and orcinol in high molar HCl

was monitored at 660nm. Results showed that the lowest concentrations of pentosans

were present 24 hours after bacterial inoculation in both stress and control media. Low

levels in the stress media were 0.011mg/ml and 0.012mg/ml in the control cultures spent

fluid. This suggests that the highest concentrations of non pentose carbohydrates would

occur around this time since results form the Dubois assay show a stable concentration of

carbohydrates in the media. After 24 hours, pentose concentration increased slightly in

the peroxide cultures back up to levels 0.019mg/ml after 120 hours. In the control media,

pentose concentrations increased more rapidly than in the stress media, with pentose

concentrations rising to 0.02mg/ml after 48 hours and peaking at 0.022mg/ml at the 96

hour point. The pentose concentrations after 24 hours clearly were not sufficient to

account for the carbohydrate levels found in the Dubois assay, meaning that there must

be substantial hexose production in the media of both control and stress cultures.

Gas Chromatography

Upon realizing that there must be significant production of hexose sugars in the

media of both xylose cultures, spent fluid samples were sent away to Paprican labs in

Montreal for carbohydrate analysis. Results indicated significant glucose production in

both control and stress samples, as well as the production of some mannose. In the

control fractions, samples contained; 244mg/L xylose, 388mg/L mannose and 894mg/L

of glucose. This is a considerable amount of hexose production given that initial xylose

levels in the media were 3g/L. This equates to a 29% conversion of xylose to glucose

and a 10% conversion to mannose in the control cultures. In the stress cultures, samples

contained, 275mg/L xylose, 427mg/L mannose and 568mg/L of glucose. Therefore,

more conversion of xylose to mannose occurred in the stress cultures and less conversion

to glucose. The conversion of xylose to glucose in the stress fractions was 18% and for

mannose 14 %. Thus, the control media appeared to be much more efficient at producing

glucose, while being slightly less efficient at producing mannose. The actual

concentrations received from the gas chromatography results differed when compared

with those from both the Dubois and Bial’s assay. However, the assay results are still

helpful when trying to understand relative levels of carbohydrates along various isolation

time points. The efficiency of glucose production, especially in control cultures, was

quite significant and could be a sufficient driving force with regards to the economic

viability of this process. Further trials would be beneficial in trying to elucidate even

higher levels of glucose and mannose from the xylose. Given that pentose concentrations

were lowest at the 24 hour point, this would be the optimal point for hexose isolation.

Xylose Dehydrogenase Characterization

Initially a BN PAGE was performed to test if xylose dehydrogenase activity

occurred in control and stress fractions. Activity was noted in membrane fractions of

both control and stress cultures. In order to better understand the manner in which xylose

is metabolized in my cultures I performed another BN PAGE to observe the activity of

XDH between control and stress fraction. High activity of XDH would suggest xylose

was being metabolized through the Weinberg pathway for quick entry into the TCA or

entry into the gluconeogenic pathway for glucose production. If XDH levels were low

this would indicate xylose reduction into xylitol by xylose reductase and subsequent entry

into the Pentose Phosphate Pathway. I also wanted to determine the affinity of XDH for

either nadp+ or nad+. This would give insight as to whether XDH is more important in

generating nadph for anti­oxidant defense, or nadh to help balance the overall energy

budget of the cell. Cellular membrane fractions were isolated 30 hours after inoculation

when the cultures were already confluent. The activity appeared to be greater in control

fractions in all four reaction mixtures prepared. XDH demonstrated a clear preference

for nadp+ over nad+. This was illustrated by the fact that although there was a

reasonably strong activity band at nad+ levels of 0.5mM, when the cofactor concentration

was reduced to 0.1 mM there was little to no XDH activity. On the other hand, XDH

produced strong band at both concentration of nadp+ with the reaction mixture containing

the 0.5mM NADP+ showing the most substantial activity bands.

The above results which illustrate that nadp+ dependant XDH is active in both

control and stress cultures shows that the Weinberg pathway is being used predominantly

to metabolize xylose. Strong activity bands in the control fractions indicate that the

nadp+ dependant pathway has a significant role in xylose degredation. Upregulation of

nadp+ dependant XDH would suggest that ATP levels in the cell are being achieved

through the quick entry into the TCA cycle provided by the Weinberg pathway. It is clear

however that much of the pyruvate pool created by the degredation of xylose is being

routed though gluconeogenesis in order to build up glucose for storage. XDH bands in

peroxide fractions were not as strong as in control fractions. The time to confluency and

protein concentrations at confluency for the two cultures were quite similar. This

indicates that in the peroxide cultures, xylose was being metabolized through a

combination of the Weinberg pathway as well as through the xylose reductase mediated

pathway. Peroxide has been shown to increase the production of ROS, and consequently

induce ETC disfunction as well as inhibit key TCA enzymes.(Mailloux et al).

Xylose Reductase Pathway Weinberg Pathway

Figure 11 Metabolic Pathways for D­Xylose in Soil Bacteria

In peroxide stressed cells it is likely that xylose was being metabolized into

mannose as a means to produce energy anearobically to compensate for lowered aerobic

ATP production. This could easily occur through the transformation of ribulose 5

phosphate, a PPP intermediate, into fructose 5 phosphate. Fructose 5 phosphate could

then be isomerized into mannose 6 phosphate and subsequently transformed into

mannose coupled with the production of one mole of ATP by mannose kinase. This

appears to have been occurring as higher production of mannose in stressed cells

illustrates up regulation of mannose kinase activity. Xylose metabolism into fructose 6

Citric Acid Cycle

Xylotal Lactone

Nadp+ Nadph

pyruvate

? Xylose reductase

XDH

Alpha Keto­Glutarate

Gluconeogenesis

Citric Acid Cycle

Xylotal Lactone

Nadp+ Nadph

pyruvate

? Xylose reductase

XDH

Alpha Keto­Glutarate

Gluconeogenesis

phosphate quickly through xylose reductase and the PPP would account for down

regulation of XDH and the weaker activity bands observed in peroxide fractions.

Optimization and Future Research

In order to try and further increase the amount of xylose being converted into

glucose two more 20mM xylose cultures were started. Low phosphate and 0.02mM Zinc

cultures were started. These cultures were both hypothesized to shut down the TCA

cycle. Zinc was used to try to knock out TCA enzymes, whereas low phosphate cultures

should have limited the proliferation of the bacteria. Both of the above processes should

have limited proliferation of the bacteria and steered the metabolism in the cultures

toward the production of storage polymers, hopefully glucose. The growth profiles of the

zinc and low Phosphate cultures were similar to the control and peroxide cultures in their

biomass at confluency; however confluency did not occur till 48 hours post inoculation.

There was limited time to analyze the metabolites in the cultures of these bacteria and

only a Bial’s test was performed. Pentose concentrations in the media after confluency

were even lower than in the control and peroxide stressed cultures and it is possible that

slightly more glucose was produced. Without performing gas chromatography or HPLC

analysis it is impossible to stipulate as to what metabolites were being produced and how

large their volumes.

Future steps forward for this project would involve studying more closely the

metabolites in the zinc and low phosphate cultures. However, a slower time to

confluency is a slight hindrance in their feasibility as cultures could only be refreshed

half as often as in the control and peroxide media. It appears that the control culture was

the most efficient at producing glucose. By growing these cultures and large media

cyclers one could refresh the media and isolate glucose and mannose every 24 hours.

The glucose and mannose are both valuable products, and large amounts of glucose could

be further fermented into ethanol or butanol for use as biofuels. Further enzyme analysis

should be performed on key metabolic enzymes such as pyruvate dehydrogenase. The

understanding of enzyme regulation at key points would allow for even better growth

media to be designed for maximum production of value added products.

In conclusion it is clear that nutrient engineering can be a useful tool in guiding

the metabolism of microbes into producing value added products. The project uncovered

a novel method for transforming xylose into More research and practice needs to go into

understanding the effects of various media on metabolic enzymes in order to maximize

the efficiency of these processes.

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