Hawkes P.W., Spence J.C.H. (Eds.) Science of Microscopy. V.1 and 2

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1050 M. Amrein

For high-quality optical images, the objective lenses are optimized

for thin glass coverslips, which are mechanically unstable sample sup-

ports for AFM imaging. The working distance of high-magniﬁ cation

objectives is very small, so the objective lens must be positioned close

beneath the sample, often with an oil or water droplet between the lens

and the coverslip. These considerations must be addressed by appro-

priate design of sample holders (which must generally also function as

a liquid cell) that are able to provide enough mechanical support for

the sample without interfering with access for the microscope objective

beneath.

For common biological samples such as cells or vesicles, there is very

low optical contrast in brightﬁ eld illumination, so some enhanced

optical contrast mechanism is required. The most common choices are

phase contrast (which provides contrast based on the sample refractive

index) or differential interference contrast (DIC, which is sensitive to

local differences in the refractive index from point to point). In both

cases, the path length and optical quality of any components in the

illumination path are critical. To use the optical microscope in its stan-

dard conﬁ guration, and hence have the optimal optical system, the

microscope requires illumination from above, using the condensor

optics. This means that the illumination light must pass “through” the

AFM in some way. The ideal optical solution is to be able to use the

condensor optics provided by the optical microscope manufacturer,

and build the AFM on an open frame around the optical path that does

not interfere with the sample illumination.

Another issue for experiments involving simultaneous AFM and

optical microscopy is the laser illumination used to measure the can-

tilever deﬂ ection. Although the majority of this is reﬂ ected by the

cantilever, a signiﬁ cant proportion spills over the cantilever edges and

can pass through the sample into the optical microscope image. For

brightﬁ eld, phase contrast, or DIC this can be removed using a simple

ﬁ lter in the optical path to cut the required wavelength, but for ﬂ uo-

rescence this can severely limit the wavelength regions available and

hence dyes that can be used for labeling. This is particularly a problem

when the sample should be labeled with more than one dye to identify

different components, or in more advanced optical techniques such as

FRET, where two ﬂ uorescent dyes are used simultaneously. Typically

most AFMs used a detection laser in the red region of the optical spec-

trum, because of the availability of small, low-power red laser diodes.

More recently, AFMs are becoming available with infrared laser illu-

mination, which allows the full visible spectrum to be used for optical

imaging. Using an infrared detection laser also results in better per-

formance of the AFM itself, in that it removes artifacts caused by

optical interference between the light reﬂ ected from the cantilever and

from the sample in the path to the photodiode detector.

2.7.2 Scanners

Over time, the scan range available from commercial AFMs has

increased, particularly the z-range or distance the tip is able to move

up and down over the surface. For cell imaging, a z-range of at least

Chapter 16 Atomic Force Microscopy in the Life Sciences 1051

10–15 µm is required, otherwise the tip will be unable to move high

enough to scan over the cell nucleus. Larger x and y scan ranges of

100 µm or more are also generally required for cell imaging, and some-

times also for imaging much smaller samples, which may be distrib-

uted inhomogeneously over the surface. The piezoceramic material

used for all AFM scanners suffers from various problems of nonlinear-

ity, hysteresis, and creep. Although it is possible to move the tip very

precisely, this is against a large background of position changes due to

longer term effects in the piezo material, as it continues moving slowly

for a long time after a voltage jump is applied (creep) or moves variable

distances depending on its history (hysteresis). These problems have

been addressed by adding position sensors (such as capacitive or

strain-based sensors) along the movement axes, so that the tip move-

ment is no longer set merely by converting the desired position into a

simple voltage. The current position of the tip is constantly read by the

position sensors and the nonlinearity of the piezo material and its

changes over time can be constantly corrected, using another feedback

loop. These “linearized” piezo systems are also becoming available in

commercial AFM systems and are likely to become standard.

There are two possibilites generally used for the lateral scanning for

an AFM—tip scanning or sample scanning. For general AFM, these

are equivalent, and both have their advantages and disadvantages, but

in terms of optimizing an AFM for biological samples, tip scanning

generally has clear advantages. On a basic practical level, much of AFM

imaging for life science research has to take place in aqueous solutions,

usually containing a reasonably high concentration of salt, and there

is always a greater risk of expensive damage when the high-voltage

piezoelectronics sit at a lower level than the sample. Even using a tip-

scanner, the AFM must be carefully designed so that the electronics

and piezo elements above the sample are protected from accidental

spillage or water vapor. This is particularly important if the sample is

to be heated to 37°C, when the evaporation is signiﬁ cant, and con-

densed water vapor could easily collect in an unsealed AFM head.

There is also a more fundamental problem with sample scanning,

however, as simultaneous AFM and optical imaging cannot generally

be performed. When the sample is moved in the z direction, which

usually lies along the optical axis, the objective lens can be tracked

with the sample using a piezoactuated lens holder, but this is not pos-

sible for lateral scanning movements across the optical axis.

2.7.3 Sample Environment

In addition to the considerations already discussed about the need for

using coverslips as a sample support for many optical imaging applica-

tions, there are other practical issues raised by life science samples.

Temperature control of the sample is often important for studying

molecular reactions or whole cells under physiological conditions, and,

for example, many lipid bilayers used as model cell membranes undergo

phase transitions over the range from around 10 to 37°C and above.

Perfusion is also important, particularly for in situ experiments and to

introduce molecules for reactions, blocking, or to change the properties

1052 M. Amrein

of the solution. For live cell work, it may also be necessary to allow a

gas exchange to equilibrate the cell medium with 5% CO

The priorities for the design of the cantilever and sample holders

should also include the easy disassembly and cleaning of all the com-

ponents that are in contact with the sample, so that, for example, ultra-

sound or autoclaving is possible. When working in liquid, all parts of

the instrument that come in contact with the ﬂ uid are sources of pos-

sible sample contamination. The issues are somewhat different depend-

ing on the applications. For instance, if the aim is to image or stimulate

living cells, then molecular-sized contamination is not so important for

AFM imaging itself, but the sample must remain sterile and the cells

must not be exposed to any chemical contaminants that will produce

a biological response. For single molecule imaging, the molecules may

not be affected by traces of certain ions leaching from metal surfaces,

but every macromolecular contaminant that could adhere to the surface

is a problem for AFM imaging.

3 Sample Preparation

To study their native structures and probe their functions, most cellu-

lar or macromolecular samples need to be kept in an aqueous environ-

ment. Hydrophilic and hydrophobic interactions promote correct

folding of the polypeptide chains into a protein and are responsible for

the formation of micelles, bilayers, and membranes from lipids and

proteins. The conformation of membrane proteins is determined by

hydrophobic interactions with lipidic tails and hydrophilic interactions

with their heads and the surrounding water (Haltia and Freire, 1995;

Engel et al., 1992; Jap et al., 1992). The pH, electrolyte type, and its

concentration and temperature also inﬂ uence the structure and func-

tion. The function of macromolecular structures depends not only on

their native conformation but often requires even more exacting envi-

ronmental conditions with respect to pH and temperature and may

depend on the presence of coenzymes or ATP, for example. Living cells

are sensitive not only to pH, ionic strength, temperature, and CO

levels; they usually need a speciﬁ c and often highly complex medium

in which to grow. When biological structures are allowed to air dry,

they are subjected to a high force caused by the change in surface

tension as the water evaporates. The energy involved is considerable.

As a result, even macromolecules become severely ﬂ attened and col-

lapsed (Baumeister et al., 1986; Kellenberger et al., 1982; Kellenberger

and Kistler, 1979; Wildhaber et al., 1985). For these reasons, most sample

preparation techniques described below are for samples in buffer. Most

of the time, immobilizing cells and macromolecular structures is all

that is needed for AFM analysis. AFM analysis of macromolecular

samples is demanding with respect to the cleanliness of the support

and the purity of the buffer. All surfaces become immediately covered

with hydrocarbons when exposed to ambient air. Even double-distilled

water can be a source of organic contaminants. A layer of these hydro-

carbons on the sample or the probe can be most disturbing for AFM.

Chapter 16 Atomic Force Microscopy in the Life Sciences 1053

As a result, the sample supports should be prepared or activated imme-

diately before use. Ultrapure water (fresh milli-Q water; ≤18 MΩ cm

−1

)

should be used to prepare all buffer and rinsing solutions because it

contains fewer hydrocarbons and macroscopic contaminants than con-

ventional bidistilled water.

In the following sections, we will describe suitable supports and

immobilization techniques for both macromolecular and cellular

samples.

3.1 Macromolecular Samples

Immobilizing macromolecular structures aims for a homogeneous dis-

tribution of the specimens in a close-to-native conformation. Tight

binding of the biological specimens to the support surface will prevent

them from clustering. They may also better withstand the forces that

arise between the probe and the sample for most AFM. On the other

hand, the structure can be substantially distorted and proteins may

even denature by strong binding. This is well known from transmis-

sion electron microscopy (TEM) as well as from STM of biological

specimens (Baumeister et al., 1986; Wang et al., 1990). The specimens

may adsorb with preferential orientations depending on the binding

conditions (Fisher et al., 1978; Hayward et al., 1978; Karrasch et al., 1993;

Müller et al., 1996). In addition to suitable binding properties, the

support should be as smooth as possible so that it does not interfere

with the structure of the biological specimen in the ﬁ nal image. Fur-

thermore, it should be relatively chemically inert to prevent contamina-

tion due to the solution or nonspeciﬁ c reactions with the biological

system.

3.1.1 Specimen Supports

Glass coverslips are widely used as an amorphous specimen support

and can be used either unmodiﬁ ed or altered to change their physi-

sorption or chemisorption properties. The surface can be almost

featureless on the scale of macromolecular specimens. They are best

suited for all experiments in which visible light is transmitted across

the sample, as in SNOM or in the combined light and atomic force

microscopy. Before use, organic contaminants, dust, and other particles

are removed by washing one time with concentrated HCl/HNO

(3 : 1)

and ﬁ ve times for 1 min with Millipore water in an ultrasonic bath

(50 kHz). This process makes the coverslips clean and smooth (rms

roughness ∼0.5 nm).

The most commonly used support for imaging biological specimens

in the AFM is mica. Mica minerals are characterized by their layered

crystal structure. Mica can be readily cleaved for a clean, atomically

ﬂ at surface. Muscovite mica (Mica New York Corporation, New York,

NY) is the most commonly used form. The average surface charge

density of muscovite mica in water is σ

≈ −0.0025 C/m

(0.015 electron

per surface unit cell).

Gold surfaces can be easily prepared by vapor deposition. They are

chemically inert against O

and stable against radicals. They bind

organic thiols or bifunctional disulﬁ des with high afﬁ nity, which can

1054 M. Amrein

be used to covalently attach biological macromolecules (see below).

Hegner et al. (1993) developed a relatively simple and reliable method

for preparing ultraﬂ at gold (Knebel et al., 1997) surfaces. They consist

of atomically ﬂ at terraces many microns in diameter. Thin carbon ﬁ lms

commonly used in TEM are smooth on molecular dimensions and

adsorb macromolecules well when freshly prepared. High-vacuum

carbon evaporators are common in electron microscopy laboratories.

3.1.2 Physiorption, DLVO Force

The most common technique for immobilizing biomolecules onto a

support is by physisorption. Usually, the objects are immobilized out

of an aqueous solution. They become attached to a support when there

is an overall attractive force that pulls the surfaces into contact. The

relevant force for adsorption is the DLVO force. Hydrophobic and

hydrophilic interactions may also play a role. The DLVO force for two

planar surfaces is (Israelachvili, 1991)

FFzFz e

DLVO el vdW

su sp

per unit area=

()

−

σσ

εε π

;

The DLVO force between charged surfaces is highly susceptible to ion

concentration and conditions can thus be adjusted to achieve good

adsorption (Müller et al., 1997) (Figure 16–19).

When the electrical double layer repulsion between the two sur-

faces has “vanished,” they rapidly coalesce to minimize the interaction

energy (Israelachvili, 1991). For the example shown in Figure 16–19, at

electrolyte concentrations above 50 mM KCl, the amount of adsorbed

membranes rapidly increased and reached its maximum at about

150 mM KCl. Note that adsorption occurred even though both mica

and the sample carried a net negative charge.

0.001 M

0.005 M

0.01 M

0.05 M

0.10 M

0.15 M

1 M

-5 10

-13

10 10

-13

5 10

-13

-10 10

24681012

[

]

F[N/nm

]

total

Figure 16–19. Dependence of the DLVO force on ion concentration (1 : 1 mon-

ovalent electrolyte) and distance between a macromolecular sample (purple

membrane) and a mica support. (From Mueller et al., 1997a, reprinted with

permission.) Whereas the attractive van der Waals force is mainly unaffected

by the electrolyte, the double layer repulsion decreases with increasing salt

concentration. The surface charge densities were −0.0025 C/m

for mica

(Israelachvili, 1991) and −0.05 C/m

for purple membrane (Butt, 1992), respec-

tively. The Hamaker constant was 3 × 10

−19

Chapter 16 Atomic Force Microscopy in the Life Sciences 1055

3.1.3 Physisorption, Hydrophobic and Hydrophilic Interaction

There is an attractive interaction between hydrophobic surfaces in

water. The attractive interaction potential is larger than the van der

Waals potential and can be very long range. The nature of these long-

range forces is not yet fully elucidated. Hydrophilic molecules, on the

other hand, tend to disorder the surrounding water molecules and

prefer contact with water molecules. Hence, the molecules repel each

other. These repulsive, hydrophilic forces are also referred to as hydra-

tion, structural, or solvation forces. They may cause the DLVO theory

to fail at small distances between two hydrophilic surfaces. With

respect to adsorption, hydrophobic molecules do not attach to a hydro-

philic surface and vice versa. For example, hydrophilic purple mem-

branes did not adsorb to highly hydrophobic supports such as

derivatized glass. The hydrophilic and hydrophobic interaction can

cause an oriented adsorption of molecular structures.

3.1.4 Physisorption, Preparation of the Support

With mica, an active surface is conveniently obtained by cleaving the

layered mica crystals prior to specimen adsorption. For most other

supports, the active surface cannot be produced so easily. These sup-

ports are usually covered by hydrocarbon contaminants and behave

more or less hydrophobicly. Glass, silicon wafers, and many thin ﬁ lms

can be rendered hydrophilic by exposure to glow discharge (for

example, in a Harrick Plasma cleaner, 1 min, p = 0.1 m bar, with air as

the residual gas) right before use. Thin carbon ﬁ lms become negatively

charged. For those specimens that adsorb better to hydrophobic sur-

faces, glow discharge must be omitted.

Coating is another way to improve physisorption on many specimen

supports and it has been used for a long time by electron microscopists

(Jacobson and Branton, 1977; Mazia et al., 1975). For example, poly-l-

lysine can be used for coating glass and mica and render the coated

surfaces positively charged. This allows cells, tissues, and plasma

membranes that are usually negatively charged to be readily adsorbed.

Objects that carry charge in an uneven distribution can be adsorbed

in a deﬁ ned orientation on a poly-l-lysine-coated surface. For example,

purple membrane mainly consists of a light-driven proton pump that

builds up an electrochemical potential across the membrane. Illumi-

nated by light, purple membranes show an asymmetric charge distri-

bution and adsorb to polylysine-coated surfaces in an oriented fashion

(Fisher et al., 1977, 1978; Hayward et al., 1978). More than 90% of the

membranes attach with their cytoplasmic surface toward the poly-l-

lysine under speciﬁ c conditions (pH 9). At a pH below 4, the majority

of the membranes (>94%) were directed with their extracellular surface

toward the coated surface.

3.1.5 Chemical Bonding

Covalent bonding can be a very reliable technique to allow ﬁ rm binding

of biological specimens to a support. Some of the ﬁ rst high-resolution

AFM topographies of protein structures in buffer solution have been

obtained using this technique (Karrasch et al., 1993). It appears that

covalent binding does not interfere with the macromolecular structure

1056 M. Amrein

anymore than physisorption. Bonding of the macromolecular spe-

cimens can be accomplished using chemically modiﬁ ed supports.

Karrasch et al. (1993) developed a protocol to cross-link biological

systems to a silanized glass coverslip. The silane (APTES, Fluka Chemie

AG, Buchs, Switzerland) contained a free amino group that allowed it

to react with the succinimide ester group of the photocrosslinker ANB-

NOS (Fluka Chemie; λ = 312 nm). Proteins were then bound to the

interface by activating the photocrosslinker with UV radiation. This

method resulted in the ﬁ rst high-resolution images of protein struc-

tures by AFM in buffer (Figure 16–20).

Epitaxial gold surfaces can effectively be functionalized by alkane-

thiols. They form ordered, self-assembled monolayers that are tightly

bound to the gold surface via chemisorption of the sulfur atoms. The

monolayers are further stabilized by the lateral hydrophobic interac-

tions of the alkyl chains (Hegner et al., 1993; Sellers et al., 1993; Wagner

et al., 1994; Wolf et al., 1995). The latter can carry head groups at the

free end that allow oriented covalent anchoring of macromolecular

structures (Allison and Thundat, 1993; Hegner et al., 1993, 1996; Wagner

et al., 1994). Wagner et al. (1995, 1996) bound protein structures via their

amino groups with an N-hydroxysuccinimide-terminated monolayer

on gold.

3.1.6 Langmuir–Blodgett Films

There are amphiphilic substances that naturally form insoluble mono-

molecular ﬁ lms on an air–water interface. They exhibit a water-soluble

polar or charged head group and a highly apolar tail. This causes them

to attach to an air–water interface with the head group immersed in

the water and the tail toward the air. The most prominent example is

the pulmonary surfactant that forms at the interface of the respiratory

Figure 16–20. Hexagonally packed intermediate layer. Scale bar = 70 nm.

(From Karrasch et al., 1993, reprinted with permission.)

Chapter 16 Atomic Force Microscopy in the Life Sciences 1057

gas lumen and the solvation layer that covers the alveolar epithelium

of lungs. Surfactant layers can be formed ex vivo in a Langmuir trough

to study their biophysical properties under deﬁ ned conditions or for

the purpose of microscopic examination. Langmuir ﬁ lms of lipids have

also been used to mimic biological membranes (for references see

Bader et al., 1984), or they served as a substrate to bind and crystallize

proteins in two dimensions for TEM and AFM investigations (e.g.,

Brisson et al., 1994). AFM proved to be outstandingly well suited to

study the structure and mechanical properties of these thin layers.

To prepare ﬁ lms for microscopy, the amphiphilic substances are

spread at the air–water interface of a Langmuir trough. They are then

compressed by a movable barrier by a desired amount. To perform

AFM on the air side of the ﬁ lm, the monolayers may be transferred

from the air–water interface onto a solid support by slowly pulling

a hydrophilic support out of the aqueous phase across the interface

(Langmuir–Blodgett transfer; Blodgett and Langmuir, 1937). The ﬁ lm

is deposited as the support is moved vertically across the air–water

interface. It is then inspected by AFM in air. To do microscopy on the

aqueous side of the ﬁ lm, the monolayer may also be deposited by

dipping a hydrophobic substrate from the air side across the interface

into the water. If a ﬁ rst lipid layer is deposited on the upstroke onto a

hydrophilic substrate and then another layer added on the down stroke

of the sample, a complete lipid bilayer has formed. This bilayer may

contain membrane proteins. It is interesting to note that deposition of

a bilayer onto a mica substrate arrests the lipid of the ﬁ rst lipid layer.

These lipids are no longer free to diffuse in the plane of the membrane.

If the support is glass, both the lipids bound to the support and those

within the second layer facing the aqueous phase are free to diffuse.

Finally, Langmuir–Blodgett transfer may not be necessary and ﬁ lms

of pulmonary surfactant have been studied directly at the air–water

interface (Knebel et al., 2002).

3.2 Cells

Successful immobilization of living cells largely depends on the cell

type. There are cells with adherent growth (for example, epithelial

cells, ﬁ broblasts, or glial cells), and cells that grow in suspension

without contact to a substrate (for example, bacterial cells or erythro-

cytes). Adhesive cells are more readily imaged with the AFM, whereas

cells that grow in suspension have to be immobilized to be imaged. It

is notable that cells may change their shape, physiology, and even their

life cycle once bound to a substrate. A variety of techniques have been

developed to immobilize living cells. Cells are best imaged with an

AFM that is combined with a light microscope.

3.2.1 Adsorbing Cells on Glass Coverslips

Cells that naturally adhere to a substrate can either be cultured on an

appropriate support and subsequently imaged, or plated on the sup port

and monitored shortly after they have established cell–substrate contact.

For both procedures, the glass coverslip must be thoroughly cleaned. If

cleaned with water, the glass support has to be dried in air or a stream

1058 M. Amrein

of N

to prevent plated cells from possible osmotic shock. Coverslips

have been coated with poly-l-lysine, collagen (Henderson et al., 1992),

proteoglycans, laminin, or ﬁ bronectin to improve adhesion.

For imaging individual, adherent cells with the SFM, the density of

the cell suspension has to be chosen such that enough space remains

for the cells to spread out. The time required for the cells to attach and

spread depends on the cell type. Before imaging, the samples have to

be rinsed with buffer solution to remove cells that are not ﬁ rmly

attached and, if feasible, examined by conventional light microscopy.

Speciﬁ c cells that were cultured on a solid support spread out to a

thickness of less than 100 nm over large areas in the periphery (Fritz

et al., 1994; Henderson et al., 1992; Hoh and Schoenenberger, 1994;

Kasas and Ikai, 1995). In these thin regions it is possible to monitor the

organization of the intracellular cytoskeleton (Figure 16–13).

3.2.2 Immobilizing Nonadhering Cells

A stable immobilization of cells that grow in suspension and do not

establish substrate interactions in their natural environment is difﬁ cult

to obtain. Hörber et al. (1992) have a method to trap single cells by a

micropipette and image the exposed part with the AFM. The setup

makes it possible to use the advantages of the micropipette technique

and to enhance the inner pressure of the cell. This is an advantage,

because the “spring constant” of a cell surface may be very low [for

example, ∼0.002 N/m (Hoh and Schoenenberger, 1994)]. Hence, the cell

is extensively deformed by any reasonable interaction force with an

AFM probe.

Permeable supports provide the possibility of measuring additional

properties of cells (for example, permeability, diffusion, and voltage

characteristics of the plasma membrane) while they are imaged by

AFM. The cells may attach onto substrates with a much smaller pore

size than the average diameter of the cell, or individual cells may be

trapped in pores that are only slightly smaller than the average cell

diameter. Kasas and Ikai (1995) have used Millipore ﬁ lters (Millipore

PCF, Millipore Corp., Bedford, MA) with pore sizes similar to that of

the cell diameter for trapping yeast cells. Hoh and Schoenenberger

(1994) have cultured MDCK epithelial cells (average lateral diameter

∼10 µm) on polycarbonate ﬁ lter supports (Millipore PCF, 12 mm diam-

eter) with a much smaller pore size (0.4 µm) than the average cell

diameter.

4 Imaging and Locally Probing Macromolecular and

Cellular Samples: Examples

4.1 Imaging

By controlling the ionic strength and composition of the deposition

buffer, individual collagen molecules can be assembled and adsorbed

onto a mica surface in various different conformations (Jiang et al., 2004).

Around ﬁ ve collagen molecules associate form microﬁ brils, which have

a lateral size of around 3–5 nm. These microﬁ brils are also likely to be an

Chapter 16 Atomic Force Microscopy in the Life Sciences 1059

intermediate stage in the formation of the larger collagen ﬁ bers seen in

natural tissue. A monolayer of these microﬁ brils can then be adsorbed to

the surface to form a nanostructured, biologically active surface. The

ﬁ brils can be aligned through adsorbing under conditions of hydrody-

namic ﬂ ow (Figure 16–21); Figure 16–21 shows a case in which the ﬁ brils

are adsorbed at very low coverage, and the 67-nm banding repeat can be

seen along the ﬁ lament length. For samples in which a complete mono-

layer is formed, the composition of the adsorption buffer can be used to

control whether the ﬁ laments organize themselves such that the bands

on adjacent ﬁ laments are aligned or not (Jiang et al., 2004).

The nanometer-scale topography of these surfaces can be measured

using AFM, and then used for cultivating cells in situ. The orientation

and growth direction of ﬁ broblast cells grown on the aligned collagen

supports depend critically on the alignment of the D-banding between

adjacent collagen ﬁ brils. The overall alignment of the direction of

the collagen ﬁ bers is not sufﬁ cient to produce a response in the cells,

but when the banding of the ﬁ bers is aligned, the cells show a strong

response (Poole et al., 2005). This is one case in which the ability to

combine AFM and optical microscopy allows the study of a biological

structure/function question over the size scale from the molecular

structure and organization to the response of whole cells.

4.2 Beyond Imaging

One strength of the AFM is that it is able to combine imaging

modes sensitive to different properties of the sample with direct mea-

surements of forces and interactions. These measurements can be

carried out at particular points (selected, for instance, from a sample

that has just been imaged), or built up over a grid to “map” the surface

properties.

Figure 16–21. Collagen ﬁ brils adsorbed on mica; sample courtesey of Müller;

imaging JPK Instruments, intermittent contact mode in buffer. Scan area =

2.6 × 2.3 µm, z range = 2.5 nm. The 67-nm banding along the axis of the indi-

vidual collagen ﬁ brils can be seen.