Thesis Paper

CHAPTER 1: INTRODUCTION

1.1 Coagulation

Coagulation is brought about primarily by the reduction of the repulsive potential of

electric double layer (1). The chemical coagulation of turbid and/or naturally colored

waters involves the interaction of particulates and/or colloids with a destabilizing agent.

The essential purpose of coagulation is to aggregate these particles into larger sizes that

will settle quickly within an hour or two, which will then be removed by sedimentation

(3). Several inorganic salts of iron and aluminum such as alum, lime, ferric chloride,

ferric sulfate, copper-as, sodium aluminate and aluminum chloride are available

commercially for coagulation. But the common coagulants used are alum and ferric

chloride (2, 3). These coagulants can effectively remove broad range of impurities from

water, such as colloidal particles and dissolved organic substances. There modes of action

can be generally explained in terms of two distinct mechanisms: charge neutralization of

negatively charged particles by cationic hydrolysis products and incorporation of

impurities in an amorphous hydroxide precipitate.

The Electrostatic stability of particles is of concern in coagulation of natural waters.

In natural waters, particles and microbes are predominantly negatively charged due to a

variety of negative functional groups on their surfaces. This negative charge was first

reported in 1929 (4). Hence, these particles may be stable as a result of electrical

repulsion. So if the surface of the particle has a net negative charge, and the bulk of

solution is electrically neutral, a potential difference exists between the surface and a

point at a distance in the bulk solution (5). In water treatment practice, such particles are

destabilized by the addition of salts causing their aggregation for rapid sedimentation and

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filtration. Destabilization of particles occur in two different mechanisms: 1) compression

of double layer (Diffuse Layer + Stern Layer) by charge neutralization and 2) adsorption

for inter particle bridging (6).

FIG. 1.1. Source of Zeta Potential and distribution of charges around a particle (7).

The electric double layer consists of two layers called the Stern Layer and the Diffuse

Layer. The stern layer (rigid layer) at the surface of the particle, consists of positively

charged counterions which may have originated from the particle or adsorption from the

solution. The total potential of this layer is called Nernst Potential. The second layer

called the Diffuse Layer is formed by the boundary between the stern layer and the bulk

solution. It is also known as the Gouy-Chapman Layer. The diffuse layer results from the

electrostatic attraction of oppositely charged ions to the particles, electrostatic repulsion

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of ions of the same charge as the particle and thermal or molecular diffusion acting

against the concentration gradients due to electrostatic effects (19). The potential gradient

over this layer is the zeta potential. The zeta potential has a maximum value at the

particle surface and decreases with distance from the surface. This decrease in the

potential is attributed to the type and concentration of ions in the bulk solution.

Addition of an electrolyte decreases the double layer thickness and hence the zeta

potential. The decrease in the repulsive electrostatic force allows short-range attractive

physiochemical interactions such as Van der Waals forces between the particles to

become significant (8). Such attractive forces cause aggregation of the particles resulting

in coagulation. This effect increases greatly with the valence of the electrolyte, which

acts as the counter ion. Aggregation increases dramatically when valence is raised from

one to two and from two to three, etc (2). Hence a trivalent cation such as Al3+ and Fe3+

will require smaller concentrations than divalent and monovalent ions to cause

aggregation (3, 9). Coagulation occurs with either a soluble hydrolytic species (i.e. Al3+)

or a solid compound (i.e. Al(OH)3) that carries positive charge in solution, under the

correct conditions of dosage and pH.

1.2 Particle-Particle Interaction

Gouy and Chapman mathematically developed the double layer theory, the results of

which has been presented by Verwey and Overbeek (1948). The double layer theory has

been developed considering the interaction between two particles in terms of potential

energy.

4

FIG.1.2. Particle-Particle interaction energies in electrostatically stabilized colloidal
systems (19).

In the above figure, ΨR is the repulsive potential energy and ΨA is the attractive

potential energy between the particles. When the particles approach each other, the

diffuse layers overlap causing a repulsive potential, ΨR, which increases in magnitude as

the distance of separation between the particles decreases. The magnitude of the

attractive energy of interactions, ΨA, which are primarily due to van der Waals forces

decreases as the distance of separation increases. The sum of potentials, ΨR and ΨA,

gives the net interaction energy between the particles. When the ΨR is greater than ΨA, an

energy barrier, Ψmax, exists between the particles resulting in electrostatic stabilization of

particles. When coagulants are added, the thickness of the double layer decreases with the

decay of zeta potential, resulting in a greater attractive potential, ΨA, between the

particles. This decreases the energy barrier, Ψmax, resulting in better aggregation and

precipitation of the particles.

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1.3 Hypothesis

This research is based on the hypothesis that the addition of coagulants to a

suspension of microorganisms and inorganic colloidal particles, reduces repulsive

electrostatic effects such that attractive van der Waals and/or hydrophobic interactions

become significant and particles are drawn together. In other words, coagulants changes

the balance of physiochemical interactions between colloidal particles facilitating

adhesion and the subsequent floc formation.

1.4 Research Objectives

As stated earlier, coagulation is one of the critical steps in drinking water treatment

processes that causes colloids, fine particles and microbes present in water to aggregate

and collect into larger particles which settle to the bottom of the sedimentation tank

before filtration. The particle-particle, particle-microbe and microbe-microbe interactions

are governed by physiochemical forces such as, hydrophobic, van der Waals, and

electrostatic forces, which in turn are determined by the interfacial surface properties of

the interacting entities (10). In addition, the physiochemical properties of water such as

pH and salt concentrations also play a significant role in these physiochemical

interactions (2).

Aluminum Sulfate and Ferric Chloride are considered non-toxic and hence widely

used as coagulants in drinking water industry. Both Aluminum and iron are trivalent

cations which form cationic and anionic complexes when added to water. These trivalent

cations form strong bonds with the oxygen atoms associated with water molecules

releasing hydrogen atoms in to the solution and thus forming aluminum and ferric

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hydroxide species (20). The solubility of metal hydroxide precipitate is an important

factor to be considered for improving coagulant performance. Amorphous Al(OH)3(am)

and Fe(OH)3(s) are formed when aluminum and ferric salts are added to water. When the

salt concentrations are less than the solubility product constants of such amorphous

hydroxides, soluble hydroxometal complexes are formed which results in coagulation of

turbid waters.

Traditionally, coagulation processes in drinking water industry are optimized by

direct inference such as Jar Tests, rather than direct analysis of physiochemical forces in

the system. Jar Tests are conducted to determine optimum coagulant dosages for the

removal of particulate matter. The results are based on rate of agglomeration, settleability

of the flocs and clarity of the supernatant water. Many variables such as temperature,

intensity and duration of mixing can affect the test results. As a result, aluminum and iron

levels in finished water are highly variable, reflecting a need for better optimization of

coagulation process by direct analysis of physiochemical forces in the system.

The atomic force microscope (AFM) is the best suited tool for the measurement of

interactive forces between colloidal particles in solution (11). Rapid advances in AFM

applications have facilitated the imaging of bacterial protein crystals, biofilms, and

microbial cells in physiological conditions (26, 12). Moreover the AFM has been used to

quantify and analyze physiochemical interactions between colloidal particles, most

notably bacteria (23). The AFM based methodology consists of immobilizing bacteria

directly onto the tip of standard AFM cantilevers, which are then used to perform force

measurements in physiological solutions.

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The Atomic Force Microscope was used in this study to evaluate bacteria-bacteria

interactions in varying concentrations of NaCl (Sodium Chloride), alum (Aluminum

Sulfate) and Ferric Chloride. The goal of this study was to evaluate the effect of the

aforementioned agents on electrostatic interactions and stability of bacterial particles in

solutions. Results from this study will ultimately be used to optimize current coagulation

processes in industry.

This research is an innovative nano-scale approach to understand and optimize

coagulation. Although coagulation is used to treat large volumes of water, the removal of

pathogens depends on the interfacial interactions of individual colloidal particles. From

an engineering perspective, this work may build a foundation of knowledge that will

facilitate the design of coagulation processes effective against current and future

pathogens. This research project is the application of a nano-scale experimental technique

to characterize and ultimately manipulate a large-scale environmental process.

CHAPTER 2: BACTERIAL STRAINS AND GROWTH CONDITIONS

2.1 Description of Bacterium

Bacteria are the simplest microorganisms that represents the most common and

ubiquitous organisms found in the environment. For example, about 1010 number of

bacteria can be found in a gram of soil (13). Bacteria are prokaryotes and they lack a

defined nucleus (25). Bacteria are classified as gram positive and gram negative based on

cell wall structure. The gram positive cell wall consists of the cell membrane (a single

lipid bilayers) surrounded by peptidoglycan and polysaccharides including teichoic acids.

The gram negative cell wall is more complex than the gram positive cell walls and

contains both inner as well as the outer lipid bilayer membranes. Between the two lipid

bilayers (periplasmic space), gram negative cell walls contain a thin layer of

peptidoglycan compared to the thick layer found in gram positives (Fig. 2.1 and Fig. 2.2).

The Gram Staining technique, developed by Hans Christian Gram, is used to classify

bacteria as gram positive or gram negative. Since gram positive bacteria have a relatively

thick peptidoglycan layer, they retain crystal violet color. whereas gram negative bacteria

have a thinner peptidoglycan component and an additional phospholipid bilayer outside

the cell wall. So, they fail to retain the violet stain and appear red.

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FIG. 2.1. Gram Positive Cell Walls (14).

FIG. 2.2. Gram-Negative Cell Walls (14).

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This research focuses on the gram negative bacteria Eschericia coli (E. coli). E. coli

was used as a representative bacteria to perform this study because it is a very well

studied microbe and is used as an indicator organism in water industry. The Cell wall

structure of E. coli has three distinct layers, they are, Inner membrane, periplasmic space

and the outer membrane. As shown in Fig. 2.1, lipopolysaccharides (LPS) forms the

outermost part of the outer membrane. The LPS molecule is anchored to the outer

membrane by a hydrophobic fatty acid region known as the Lipid A. Attached to Lipid A

is ketodeoxyoctonate (KDO), a core polysaccharide region and a relatively larger O-

antigen region (Fig. 2.3). The core and the O-antigen region of the LPS primarily consists

of six and seven carbon sugars. The hydrophilic portion of the LPS, consisting of (KDO)

through the O-antigen, extends into the surrounding aquatic environment (15). The LPS

of the E. coli K-12 strains used in this study contains only the Lipid A, KDO and the core

polysaccharide. K-12 bacteria do not synthesize O-antigens.

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FIG. 2.3. Structure of Bacterial Lipopolysaccharide showing the Lipid A, Core
polysaccharide and O-antigens (16).

2.2 Bacterial Strains and Growth Conditions

E. coli was used in this study because it is the most extensively studied

microorganism. Moreover, previous AFM studies on bacterial adhesion of E. coli K-12

strains served as a basis for data analysis in this research. E. coli K-12 D21, which is a

wild type strain, was acquired from the E. coli Genetic Stock Center (Dept. Biology, Yale

University, New Haven CT USA). This strain synthesizes the core LPS molecules but no

O-antigen (17, 18). Previous AFM-based studies found LPS molecules to play an

important role in the bacteria-surface interactions (23).

Previous studies have characterized surface properties of E.coli D21, such as

hydrophobicity (contact angle measurements) and surface charge density (ζ potential

measurements). According to Ong Y. L. et al. (51), the contact angle measurement was

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found to be 19.4±3.0 and ζ potential was -28.8±1.7. Therefore E. coli D21 was very

strongly hydrophilic and negatively charged.

In this study, bacteria were grown in Luria Broth at 37oC and harvested in the mid-

exponential phase (Optical Density, OD600 0.4 to 0.6). The cells were washed twice with

phosphate buffer saline (PBS; 136 mM NaCl, 2.68 mM KCl, 10.1 mM Na2PO4, 1.37 mM

KH2PO4; pH 7.5). Cells were then stirred with 2.5%v/v gluteraldehyde solution for three

hours at 4oC to a final concentration of 0.6-0.8 mg dry cell weight/ml. Gluteraldehyde

working solution was prepared from a 25%v/v aqueous stock solution, diluted to 2.5%v/v

in PBS buffer and then purified by stirring with 50 mg/ml charcoal at 4oC for 24hrs. The

pH of the gluteraldehyde solution was maintained at 7.5 before the addition of charcoal.

After treating the cells with gluteraldehyde, they were rinsed at least 10-15 times in 1mM

Tris buffer [tris(hydroxymethyl)aminomethane], pH 7.5 and stored at 4oC.

2.3 Bacteria Immobilization Protocol

Gluteraldehyde treated bacteria were irreversibly immobilized onto a planar glass

substrate as well as on the tip of AFM cantilever according to the protocol developed by

Razatos et al. (23). To immobilize bacteria on the planar glass substrate, the planar glass

were first cut into 1x1 cm slides, soaked in 1M HNO3 overnight, rinsed with ddH2O

followed by MeOH for a period of 30 minutes each, and dried with sterile air. Clean glass

was then coated with a positively charged polymer, 1%v/v polyethyleneimmine, (PEI;

MW = 1,200; Polysciences Inc., Warrington, PA USA) and allowed to adsorb for 2 – 3

hrs. The glass slides coated with 1%v/v PEI were then rinsed in distilled de-ionized water

(ddH2O) and stored at 4oC. 4%v/v PEI was used to coat AFM cantilevers. Both 4%v/v

13

and 1%v/v PEI were prepared by diluting distilled de-ionized water (ddH2O) and pH was

adjusted to 8 with HCl. The negatively charged bacterial cells adhere the positively

charged PEI, causing their immobilization on PEI coated cantilever tip as well as on PEI

coated glass substrate.

To immobilize bacteria onto the planar glass substrate, glutaraldehyde-treated cells in

1mM Tris were placed on PEI-coated glass substrates and incubated at room temperature

for 12 – 15 hrs.

To immobilize bacteria onto the tip of AFM cantilevers, gluteraldehyde-treated cells

in 1mM Tris were manually deposited on PEI-coated tips of AFM cantilevers. The

cantilevers were dried for 2 – 3hrs at room temperatures. Bacteria coated cantilever were

then rinsed in ddH2O and air dried.

CHAPTER 3: CRYPTOSPORIDIUM AND ITS IMMOBILIZATION PROTOCOL

3.1 Cryptosporidium

Cryptosporidium is a coccidian protozoan parasite known to infect humans. For

several decades, cryptosporidium was thought to be a rare, opportunistic animal

pathogen. However, this belief has changed over the past 20 years as cryptosporidium has

been designated as a large asymptomatic infectious parasite and an important cause of

enterocolitis and diarrhea in humans (45, 46). It is ubiquitous in the environment as per

the reports of surface water contamination all across the world (47, 48).

The parasite is protected by an outer shell that allows it to survive outside the body

for long periods of time. It infects humans, but has also been shown to infect animals like

dogs, cattle, pigs and sheep. Cryptosporidium is classified as a sporozoa, since its oocyst

releases four sporozoites upon ecxystation. Its life cycle includes both sexual and asexual

cycles (Fig. 3.1). They are;

Excystation of the orally ingested oocyst and release of the sporozoites.

Sporozoites attach to the walls of the intestine and initiate asexual intracellular

multiplication.

Production of merozoites which are again able to infect intestinal cells.

Initiation of sexual replication and fertilization causing development of the

oocysts, and

Development of infectious sporozoites within the oocysts.

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FIG. 3.1. Cryptosporidium life cycle (49).

Oocysts have been found in lakes and reservoirs, raw and treated surface waters. It

can be transmitted from fecally contaminated food and water, from animal-person contact

or person-person contact. Several water borne outbreaks of cryptosporidiosis have been

reported in recent years, most notably of which was the outbreak in Milwaukee, USA,

where up to 400 000 people were infected and 100 deaths of immunocompromised

individuals were attributed to the disease (50). No safe and effective therapy for

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cryptosporidiosis has been successfully developed. Oocysts are resistant to conventional

treatment processes such as chlorination for disinfection and this has focused attention on

the need for enhanced physical and chemical removal of oocysts from water supplies.

Since cryptosporidiosis is a self-limiting illness in immunocompromised individuals,

general, supportive care is the only treatment for the illness.

3.2 Oocyst Immobilization

Cryptosporidium oocysts were obtained from the sterling parasitology laboratory at

the University of Arizona, Tucson, Arizona. The oocysts were stored in antibiotic

solution (100 µg/ml penicillin and 100 µg/ml gentamicin) containing 0.01% tween 20.

The oocysts were washed atleast five times in phosphate buffer saline (PBS; 136 mM

NaCl, 2.68 mM KCl, 10.1 mM Na2PO4, 1.37 mM KH2PO4; pH 7.5). Oocysts were then

stirred with 2.5%v/v gluteraldehyde solution for three hours at 4oC. After treating the

oocysts with gluteraldehyde, they were rinsed atleast 10-15 times in 1mM Tris buffer

[tris(hydroxymethyl)aminomethane], pH 7.5 and stored at 4oC.

To immobilize Cryptosporidium oocysts on the planar glass substrate, the planar

glass were first cut into 1x1 cm slides, soaked in 1M HNO3 overnight, rinsed with ddH2O

followed by MeOH for a period of 30 minutes each, and dried with sterile air. Clean glass

was then coated with a positively charged polymer, 1%v/v polyethyleneimmine, (PEI;

MW = 1,200; Polysciences Inc., Warrington, PA USA) and allowed to adsorb for 2 – 3

hrs. The glass slides coated with 1%v/v PEI were then rinsed in distilled de-ionized water

(ddH2O) and stored at 4oC. 4%v/v PEI was used to coat AFM cantilevers. Both 4%v/v

17

and 1%v/v PEI were prepared by diluting distilled de-ionized water (ddH2O) and pH was

adjusted to 8 by adding HCl.

To immobilize Cryptosporidium onto the planar glass substrate, glutaraldehyde-

treated oocysts in 1mM Tris were placed on PEI-coated glass substrates and was

incubated at room temperature for 12 – 15 hrs.

To immobilize Cryptosporidium onto the tip of AFM cantilevers, gluteraldehyde-

treated oocysts in 1mM Tris were placed over PEI-coated tip of AFM cantilevers. The

cantilevers were dried for 2 – 3hrs at room temperatures. Cryptosporidium coated

cantilever tips were then rinsed in ddH2O and air dried.

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FIG. 3.2. AFM image of Cryptosporidium oocyst immobilized on a planar glass in
solution.

CHAPTER 4: MATERIALS AND METHODS

4.1 Bacteria Cell Immobilization

In order to use the AFM to evaluate bacteria-bacteria interactions in solution, the

bacteria must be immobilized on both the AFM cantilever tip and the planar substrate

probed by the tip. Razatos et al recently developed an immobilization protocol that

reproducibly and reliably immobilizes bacteria on the aforementioned surfaces.

Glutaraldehyde was found to be critical to the stability of bacterial lawns on AFM

cantilever tips and on planar glass slides (22, 23, 24).

Glutaraldehyde is a small molecule with two aldehyde groups separated by a flexible

chain of 3 methyl groups (HCO-(CH2)3-CHO) (28). In aqueous solutions, glutaraldehyde

polymerize through the aldehyde groups into molecules of variable size (27, Fig. 4.1). In

biological samples, the aldehyde group reacts with free amine groups (the amino

terminus or lysine residues) of proteins via a Schiff base reaction (Fig. 4.2). Each of the

two aldehyde groups in glutaraldehyde can form a Schiff base with different amino

groups of proteins resulting in cross-linking of the proteins (51).

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FIG. 4.1. (A) Different schematics of a monomeric glutaraldehyde molecule (28). (B)
Polymerization reaction of gluteraldehyde, showing an aldehyde side-chain on each unit
of the polymer (28).

FIG. 4.2. Schiff base reaction of glutaraldehyde with amino groups of proteins (28).

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As a result of this reaction, glutaraldehyde increases cell rigidity. Moreover it is

believed that the glutaraldehyde reacts with the polyethyleneimmine during the

immobilization of bacteria resulting in stable bacterial lawns. The glutaraldehyde does

not react with the lipopolysaccharide or exopolysaccharide molecules coating the

bacterial cell surface, which are devoid of amino acids. Extensive control studies found

that glutaraldehyde does not alter the physiochemical properties of the bacterial cell

surface and does not affect AFM force measurements (52).

4.2 Bacterial Growth and Immobilization Protocol

Bacteria-bacteria interactions were evaluated for E. coli K-12 D21 acquired from the

E. Coli Genetic Stock Center (Dept. Biology, Yale University, New Haven CT USA).

This is a typical gram negative bacteria that synthesizes the full core lipopolysaccharide

molecule but is devoid of exopolysaccharide and cell surface appendages. Bacteria were

grown overnight at 37oC in Luria Broth (LB). About 0.25ml of the overnight culture were

added to 25ml of LB and grown in a shaker at 37oC. The culture was grown to an optical

density (OD600) of 0.4-0.6. The 25ml suspension was then centrifuged for 10 minutes at

5,000 RPM. The supernatant was eliminated and the cells were rinsed with phosphate

buffered saline (PBS, pH 7.4). The cells were then added to a 2.5 vol% glutaraldehyde

(pH 7.5) solution to a final concentration of 3-6 mg dry cell weight/mL. This solution

was constantly stirred for 2-3 hrs at 4oC. After fixing the cells with glutaraldehyde, they

were rinsed 10-15 times and resuspended in 1mM Tris buffer (pH 7.4).

Standard Nanoprobe cantilevers (Digital Instruments, Santa Barbara, CA) were pre-

soaked in 4 vol% polyethyleneimmine (Sigma Chemical Co., St. Louis, MO USA)

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solution (pH 8.0) for 2 to 3 hours and then rinsed with ddH2O (distilled de-ionized

water). The bacterial cells suspended in Tris buffer (pH 7.4) described above were

centrifuged for 10 minutes at 5,000 RPM into a pellet. Bacteria from the pellet were then

manually transferred onto the polyethyleneimmine coated silicon nitride tips. The

bacteria coated tips were incubated at room temperatures for 10 minutes, rinsed with

ddH2O, and stored overnight at 4oC.

Glass microscope slides were cut into 1x1 cm slips, soaked in 1M HNO3 overnight

and rinsed for 30 minutes with ddH2O followed by MeOH. Clean glass slides were then

coated with 1 vol% polyethyleneimmine, which was allowed to adsorb for 2-3 hrs and

then rinsed with ddH2O. The bacterial cells suspended in Tris buffer were transferred

onto the PEI coated glass slides. The bacteria were incubated on the glass at room

temperatures for 12-15 hrs until excess liquid had evaporated resulting in a bacterial film

immobilized on the glass slide.

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FIG. 4.3. AFM image of E. coli D21 bacteria immobilized on a planar glass in
solution.

CHAPTER 5: ATOMIC FORCE MICROSCOPY

5.1 Introduction

The atomic force microscope is a primary form of scanning probe microscope (SPM),

developed by Binning, Quate, and Gerber in 1986. It is an instrument that can provide

nanometer-scale analysis to sample surface. Therefore AFM is broadly used in many

fields such as chemistry, physics, microbiology and material science. Imaging under

atomic force microscope has opened a range of novel applications in the field of

microbiology. Since 1980, AFM has been useful in the imaging of biomolecules, lipid

membranes, two dimensional protein crystals and cells (29, 30, 31). Recently AFM has

been used to visualize the surface structure of two-dimensional bacterial protein crystals,

biofilms and individual cells in physiological conditions. It was also used to evaluate

bacterial adhesion to biomaterials in physiological solutions by Razatos et al. (22, 24).

Progress has been made to measure biomolecular interactions and physical properties

of microbial surfaces using force spectroscopy (26). Spatially resolved AFM force

spectroscopy, combined with topographic imaging, is emerging as a valuable technique

to map physical heterogeneities of the cell surfaces. The capacity of the AFM to measure

surface forces as well as topography is well established, with measurements of van der

Waals, electric double layer, solvation, steric and hydrophobic forces (32, 33).

Sometimes modifications have to be made for the AFM probes with defined chemical

groups for quantitative measurements of the surface forces (34). Recently, Razatos et al.

(24), developed the AFM-based methodology to evaluate bacterial adhesion on

biomaterials in physiological solutions.

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5.2 Components of AFM

FIG. 5.1. Components of an Atomic Force Microscope (AFM).

The atomic force microscope (AFM) is a surface imaging technique, which operates

by sensing the force between a sharp tip and a sample surface (12). The different

components of an AFM are shown in Fig. 5.1. It consists of a laser arrangement, a mirror,

a piezo electric scanner, photodiode detector and a standard nano-probe cantilever.

Silicon or silicon nitride tips are integrated in the cantilever. The tips have a pyramidal

profile with a curvature radius ranging between 5 and 30nm. The sample is placed over a

sample holder on top of a piezo scanner. To image the samples, probe and the sample are

brought in contact and the surface of the sample is scanned by the probe. A laser beam

focused on the free end of a cantilever is reflected into a sensitive photodiode. This

optical lever is used to monitor the position of the cantilever. The light intensities

incident on the photodiode detector are converted into voltages and amplified by the

amplifier. The feedback amplifier is connected to a computer, where the image of the

sample is displayed.

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5.3 AFM Force Measurements

The AFM can also be used to measure interaction forces between the cantilever tip

and planar substrates. In this case the surface is advanced towards and retracted away

from the stationary cantilever. Tip deflection is monitored as a function of distance of

separation (Fig. 5.2)

3 1
Cantilever
Deflection 2
(nm) 4 5

Distance of Separation (nm)
1,2 and 3 represent approach curve
4 and 5 represent retraction curve

FIG. 5.2. AFM operation in Force Mode. (1) Distance between the cantilever and the
surface is sufficiently large. (2) Cantilever deflecting towards the substrate due to force
exerted on the cantilever by the substrate (Attraction). (3) Cantilever and the substrate
moving at the same rate in the constant compliance region. (4) Surface retracted away
from the cantilever. (5) Tip detaches from the substrate as the surface moves away from
the cantilever.

When the tip-sample interaction is negligible (i.e. when the distance of separation

between the cantilever and the substrate is sufficiently large) no force is exerted on the

cantilever by the substrate. So, the cantilever is not deflected towards or away from the

substrate (Fig. 5.2, line1). As the surface approaches the cantilever, at a critical distance

of separation the cantilever tip will deflect towards or away from the surface depending

on the presence of an attractive or repulsive interaction between them. In Fig. 5.2, line 2

is pointing downwards depicting an attractive interaction between the tip and the surface.

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This completes the approach curve reflecting the long-range, reversible, physiochemical

interactions between the sample and the probe. At this point, the cantilever and the

surface move at the same rate in the constant compliance region to achieve maximum

limit of compression of the sample surface (Fig. 5.2, line3). The substrate is then

retracted away from the AFM cantilever to measure the pull-off force which reflects the

strength of adhesion between the cantilever and surface (Fig. 5.2, lines 4 & 5). This

completes the retraction curve reflecting short-range, irreversible interactions between the

sample and the AFM cantilever tip.

5.4 AFM Data Analysis

AFM force measurement data are initially acquired and displayed in terms of tip

deflection (nm) versus relative distance of separation (nm) (Fig. 5.2). The deflection

curves were normalized so that the tip deflection is zero where there is negligible

interaction between the tip and the surface. Also the slope of the portion of the curve

where the cantilever moves with the surface to achieve maximum limit of compression of

the sample surface (constant compliance region) was set equal to the rate of piezo

displacement. The distance of separation (nm) was calculated as the sum of tip deflection

and piezo position relative to zero distance of separation (11). To convert the data from

tip deflection (nm) versus relative distance of separation (nm) to force (nN) versus

distance of separation (nm), tip deflection (nm) was multiplied by the spring constant of

the cantilever according to Hooke’s law, F = k * ∆X. F is force (nN), ∆X is the tip

deflection (nm) and k is the spring constant. The spring constant used in this study was

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0.06nN/nm for long, thin standard cantilevers as reported by the manufacturer (Digital

Instruments, Santa Barbara, CA).

5.5 Atomic Force Microscope Operation

AFM force measurements were performed using a Nanoscope III Contact Mode AFM

and standard Nanoprobe cantilevers with silicon nitride tips (Digital Instruments, Santa

Barbara, CA). The experiments were conducted in a fluid cell (Digital Instruments, Santa

Barbara, CA), in which bacteria coated cantilevers were placed. The planar glass slides

coated with bacteria were mounted on a piezo. AFM force measurements were carried

out by manually approaching the surface towards the AFM cantilever without touching

the two entities prior to the force measurement. To ensure the presence of confluent

bacterial lawns on the glass surface, AFM images of the bacteria coated substrate were

taken following every force measurement.

CHAPTER 6: RESULTS AND DISCUSSION

6.1 Experimental Results

The model organism used for initial AFM- based coagulation studies was the E. coli

K-12 strain D21. This strain is a wild type in the synthesis of core lipopolysaccharide

molecules that coat the bacterial surface and influence bacterial adhesion (23, 24). The

bacterial strain E. coli D21 was chosen because of its extensive use in previous AFM-

based studies. Zeta potential and contact angle measurements have found D21 to be

negatively charged and hydrophilic. The measured zeta potential was 28.8 ± 1.7 mV and

the contact angle measured with water was 19.3 ± 3.0o for E. coli D21 (51). This bacterial

strain has been successfully immobilized on AFM cantilevers and on planar glass

according to protocols by Razatos et al. (24).

In addition to E. coli, the AFM was also used to evaluate adhesion forces of

Cryptosporidium Parvum, a protozoan parasite that represents a series threat to

contamination of drinking water. Cryptosporidium oocysts are known to be negatively

charged from the electrophoresis experiments (32). The zeta potential measurements for

Cryptosporidium parvum oocysts typically vary from –15mV ± 10 mV to around –35 ±

10 mV (37, 38, 39, 40, 41).

6.2 Control Experiments

AFM based coagulation experiments were conducted in PBS buffer and in PBS +

NaCl to determine the role of electrostatic interactions between bacteria. The addition of

an electrolyte such as NaCl, decreases the double layer thickness and hence the zeta

potential resulting in destabilization of negatively charged particles in solution. This

decreases the repulsive electrostatic forces allowing short-range attractive

30

physiochemical interactions such as Van der Waals forces between the particles to

become significant. Fig. 6.1 is a representative plot of tip deflection versus relative

distance of separation between D21 bacteria immobilized on to the AFM cantilevers and

D21 immobilized on clean glass in PBS buffer. The black curve represents the initial

physiochemical interactions between the D21 bacteria during approach. The gray curve

represents the pull-off force between the bacteria-coated tip and the bacteria-coated

substrate during retraction. During AFM operation, as the surface advanced towards the

tip (approach), the cantilever deflected towards the surface when the relative distance of

separation between bacteria on the tip and bacteria on substrate was 30nm. This indicates

that there is an attractive force between bacteria on tip and bacteria on substrate in PBS

buffer (Fig. 6.1). During retraction, the cantilever deflected due to a strong binding force

between bacteria in PBS buffer (Fig. 6.1).

Fig. 6.2 is a representative plot of tip deflection versus relative distance of separation

between D21 bacteria immobilized on the AFM cantilevers and D21 bacteria

immobilized on clean glass in PBS buffer + 100mM NaCl. The black curve represents

initial physiochemical interactions between D21 bacteria during approach. The gray

curve represents pull-off force between bacteria-coated tip and bacteria-coated substrate

during retraction. During AFM operation, as the surface advanced towards the tip

(approach), the cantilever deflected towards the surface when the relative distance of

separation between bacteria on tip and bacteria on substrate was 70nm. In this case the

initial attraction between the D21 bacteria becomes significant over longer distances of

separation in comparison to PBS buffer alone (Fig. 6.1, black curve). The gray curve

31

represents pull-off force between bacteria due to binding. This pull-off force measured in

PBS + NaCl was greater than in PBS buffer alone (Fig. 6.2, gray curve). These results

indicate that the addition of an electrolyte such as NaCl reduces the repulsive effect of

electrostatic interactions such that attractive (hydrophobic and van der Waals) forces

dominate over longer distances of separation. Therefore, NaCl has a destabilizing effect

on colloid particles in solution.

32

15
10
Tip Deflection (nm)

5
Approach
0
Retraction
-5 0 20 40 60 80 100

-10
-15
-20
Relative Distance of Separation (nm)
-25

FIG. 6.1. Tip deflection versus relative distance of separation reflecting interaction
between bacteria on glass surface and bacteria on cantilever tip in PBS Buffer.

33

15
10
Tip Deflection (nm)

5
Approach
0
Retraction
-5 0 20 40 60 80 100

-10
-15
-20
-25 Relative Distance of Separation (nm)

FIG. 6.2. Tip deflection versus relative distance of separation reflecting interaction
between bacteria on glass surface and bacteria on cantilever tip in PBS Buffer + NaCl.

34

The deflection curves during the approach for both the experiments i.e., experiment in

PBS buffer and in PBS buffer + 100mM NaCl solution, were converted in to force

curves. These force curves are obtained by multiplying the tip deflection during approach

with the spring constant of the cantilever (5.4, Fig. 6.3). The black curve represents the

force curve for experiments in PBS only, whereas the gray curve is the force curve for

experiments in PBS + NaCl.

0.1
PBS+NaCl
0
PBS only
0 10 20 30 40 50
Force (nN)

-0.1

-0.2

-0.3

-0.4

-0.5 Relative Distance of Separation (nm)

FIG.6.3. Force curves between bacteria on glass surface and bacteria on cantilever tip
in PBS buffer and in PBS buffer + NaCl during approach portion of AFM cycle.

TABLE 6.1. Summary of AFM force measurements between E. coli bacteria.

Configurations E. coli bacteria on tip and on E. coli bacteria on tip
glass surface and on glass surface
Experiments In PBS Buffer only In PBS buffer + NaCl

Force values in nN -0.35 ± 0.06 -0.45 ± 0.02

35

For bacteria-bacteria interaction in PBS buffer, the force value was 0.35nN whereas

for bacteria-bacteria interaction in PBS buffer + 100mM NaCl, the force value was

0.45nN. The negative sign for the force values indicate downward deflection of the

cantilever due to attractive interactions between bacteria (Fig. 6.3 and Table 6.1). These

force values indicate that NaCl reduces the long-range repulsive interactions such that

attractive (hydrophobic and van der Waals) forces dominate.

6.3 Experiments with Coagulants

Different experiments were conducted to determine bacteria-bacteria interaction in

different concentrations of alum and Ferric Chloride dissolved in PBS buffer. Alum and

ferric chloride are the most common chemical coagulants used for water treatment.

During the coagulation process in water treatment, destabilization/charge neutralization

of negatively charged particles is caused by the cationic hydrolysis products of aluminum

and iron, and floc formation of impurities with amorphous hydroxide precipitate of

aluminum and iron. The optimum pH value for alum is variable from 5.4 - 8.0, whereas

for ferric chloride the pH value is variable between 4.5 – 12 (2). The typical dosage of

alum and ferric chloride used in water treatment varies between 5 – 50 mg/l and 10 – 80

mg/l respectively, for the removal of impurities including inorganic particles, pathogenic

microbes and dissolved organic matter (2, 42, 43, 44). The different concentrations of

alum and ferric chloride selected for our experiments were, 12, 18, 24mg/l and 20, 40, 60

mg/l respectively. These concentrations were used to closely model the conditions used

in municipal water treatment plants.

36

Fig. 6.4 is a representative plot of tip deflection versus relative distance of separation

between D21 bacteria immobilized on the standard AFM cantilevers and D21 bacteria

immobilized on a clean glass in different concentrations of alum in PBS Buffer. The

black curves, gray curves and the light gray curves represent the deflection curves for

bacteria-bacteria interaction in 12mg/l, 18mg/l and 24mg/l of alum in PBS buffer

respectively. Fig. 6.5 is a representative plot of tip deflection versus relative distance of

separation between D21 bacteria immobilized on the standard AFM cantilevers and D21

bacteria immobilized on a clean glass in different concentrations of ferric chloride in PBS

buffer. The black curves, gray curves and the light gray curves represent the deflection

curves for bacteria-bacteria interaction in 20mg/l, 40mg/l and 60mg/l of ferric chloride in

PBS buffer respectively. In both Fig. 6.4 & Fig. 6.5, thick curves represent the approach

curves and the dotted curves represent retraction curves.

37

Tip Deflections with 5nm offsets
(a)

(b)

(c)

0 10 20 30 40 50 60 70 80

FIG. 6.4. Plots of tip deflection versus relative distance of separation for bacteria-
bacteria interaction in different concentrations of alum in PBS buffer. Black curves, gray
curves and light gray curves are for 12mg/l, 18mg/l and 24 mg/l of alum respectively.
Tip deflections with 5nm offsets

0 10 20 30 40 50 60 70 80
Relative distance of separation (nm)

FIG. 6.5. Plots of tip deflection versus relative distance of separation for bacteria-
bacteria interaction in different concentrations of ferric chloride in PBS buffer. Black
curves, gray curves and light gray curves are for 20mg/l, 40mg/l and 60mg/l of ferric
chloride respectively.

38

For 12mg/l of alum in PBS buffer, the cantilever deflected towards the surface during

the approach cycle when the relative distance of separation between bacteria coated tip

and substrate was 35 nm. (Fig. 6.4(a)). With 18mg/l of alum in PBS buffer, the cantilever

deflected when the relative distance of separation was 45nm (Fig. 6.4(b)) and for 24mg/l

of alum in PBS buffer, the cantilever deflected when the relative distance of separation

was 55 nm (Fig. 6.4 (c)). The approach curves for all the three experiments indicate that

as the concentration of alum is increased, attractive interactions between bacterial cells

are occurring at a greater distance of separation. Even during the retraction cycle, as

shown by the dotted curves in Fig. 6.4, cantilever deflections increase due to greater

binding forces between bacteria on the tip and on the surface as a function of alum

concentration.

Experiments with ferric chloride coagulant showed similar results. With 20mg/l of

ferric chloride in PBS buffer solution, the cantilever was observed to deflect towards the

surface when the relative distance of separation between the bacteria coated tip and

bacteria coated surface was 20nm (Fig. 6.5(a)). With 40mg/l of ferric chloride in PBS

buffer, the cantilever deflected towards the surface at 25nm and with a larger tip

deflection (stronger force) compared with 20mg/l of ferric chloride. (Fig. 6.5(b)). For

60mg/l of ferric chloride, the cantilever deflected at 35nm and the deflection of the

cantilever was larger in comparison to 20 and 40mg/l of ferric chloride in PBS buffer

solution (Fig. 6.5(c)). The tip deflection during retraction also increased with the increase

in ferric chloride concentration reflecting stronger binding interactions between the

bacteria.

39

0.5 12 mg/l
0 18 mg/l
0 10 20 30 40 24 mg/l
Force (nN)

-0.5

-1

-1.5

-2

-2.5

FIG. 6.6. Representative plots of force versus relative distance of separation between
bacteria on tip of the cantilever and bacteria on surface in different concentrations of
Alum in PBS buffer.

TABLE 6.2. Summary of AFM force measurements for experiments with different
concentrations of alum in PBS buffer.

Alum Conc. in (mg/l) 12 18 24

Force in (nN) -0.78 ± 0.06 -0.81 ± 0.02 -1.98 ± 0.2

The deflection curves during the approach for all the three experiments with alum,

were converted in to force curves by multiplying the tip deflections during approach

cycle, with the cantilever spring constant. (5.4, Fig. 6.5). The negative sign indicates

downward deflection of the cantilever due to attractive interactions. For alum

concentrations of 12 and 18 mg/l, the force values were the same. But the attractive force

of interaction between bacteria more than doubled for the alum concentration of 24 mg/l.

40

The results demonstrate that alum coagulant reduces repulsive electrostatic interactions

between bacterial cells, such that attractive interactions dominate over greater distance of

separation between them. Moreover, there is a critical alum concentration that will yield

significantly stronger interaction forces between bacteria. This concentration and its

effect on interactions forces can only be elucidated using the AFM-based coagulation

methodology.

0.2 20 mg/l
0 40 mg/l
-0.2 0 10 20 30 40 60 mg/l
Force (nN)

-0.4
-0.6
-0.8
-1
-1.2
-1.4 Relative Distance of Separation (nm)

FIG. 6.7. Representative plots of force versus relative distance of separation between
bacteria on tip of the cantilever and bacteria on surface in different concentrations of
ferric chloride in PBS buffer.

TABLE 6.3. Summary of AFM force measurements for experiments with different
concentrations of ferric chloride in PBS buffer.

Ferric Chloride Conc. in mg/l 20 40 60

Force in nN -0.22 ± 0.05 -0.45 ± 0.08 -1.16 ± 0.01

41

The deflection curves during the approach for all the three experiments with ferric

chloride, were converted in to force curves (Fig. 6.7). The negative sign indicates

downward deflection of the cantilever due to attractive interactions between bacteria. For

ferric chloride concentrations of 20 mg/l, 40 mg/l and 60 mg/l, the attractive force values

were 0.22nN, 0.45nN and 1.16nN respectively, showing a progressive increase in force

as a function of ferric chloride concentration. These results demonstrate that ferric

chloride coagulant reduces repulsive electrostatic interactions between bacterial cells,

such that attractive interactions dominate over greater distance of separation between

them.

The greater attractive forces between bacterial cells may be due to the decrease in

double layer thickness and hence the zeta potential due to coagulants. The decrease in

repulsive electrostatic forces allows short-range attractive physiochemical interactions

such as Van der Waals forces between the bacterial cells to become significant. Such

attractive forces are the cause for particle aggregation and coagulation.

CHAPTER 7: SUMMARY

The AFM was used to directly measure bacteria-bacteria interactions in different

concentrations of Alum and Ferric chloride coagulants. The different concentrations of

both alum and ferric chloride used in our experiments reflect the conditions used in

municipal water treatment plants.

The bacteria used for this study was the E. coli K-12 strain, D21, which is a wild type

strain that synthesizes core Lipopolysaccharide molecules but no O-antigen. For the

purpose of force measurements, bacteria were immobilized on planar glass and on the

tips of standard AFM cantilevers. In all the experiments, the addition of coagulants

reduced the long-range stabilizing electrostatic interactions between bacteria resulting in

strong attractive interactions over longer distances of separation.

The results of this study can be summarized by the following conclusions:

Control studies (i.e., experiments in PBS buffer and in PBS buffer + NaCl,)

demonstrate that the electrostatic interactions play a dominant role in bacterial

adhesion.

Coagulants such as Aluminum Sulfate (Alum) and Ferric Chloride reduces

repulsive electrostatic interactions such that attractive force (i.e., van der

Waals or hydrophobic) become stronger over greater distances of separation

between bacterial cells.

The AFM-based coagulation Methodology makes it possible to optimize

coagulation conditions by providing quantitative data in the form of force

versus distance curves.

43

Future studies should focus on investigating specific interactions between i) microbes

that are commonly found in surface waters, other than E. coli, ii) microbes-particle

interactions and iii) particle-particle interactions in different concentrations of commonly

used coagulants in drinking water treatment processes.

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