living under an atomic force microscope

8/7/2019 Living Under an Atomic Force Microscope

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180

M. OBST AND M. DITTRICH

2005 The Authors

Journal compilation 2005 Blackwell Publishing Ltd

However, the alteration of picoplanktonic cell surfaces and

the process of mineral nucleation on those have never been

observed online or in situ

; most probably because no appro-

priate technique has been developed so far. One basic require-

ment of the technique is the capability ofin situand in vivo

observation of cells at a sufficient lateral resolution. Further-

more, long-term stability is necessary in order to be able to

perform experiments on timescales that are relevant forinvestigations on cell surface changes and mineral nucleation.

Scanning force microscopies (SFM) like atomic force microscopy

(AFM), in principle, meet the first requirement as discussed in

the following section.

Applications of AFM and scanning tunneling microscopy

(STM) in biological research are reviewed in Marti & Amrein

(1993). One decade later Jena & Hrber (2002) presented a

nice overview of more advanced techniques including in situ

AFM in fluids.

In situAFM allows imaging and characterization of topo-

graphy, physical and chemical properties of cell surfaces at the

nm-scale (Mller et al., 1999). Since in situAFM has beendeveloped and applied to biological samples, one of the most

important limitations of the previously mentioned electron

microscopy studies could be overcome: Experiments on

biological samples can be performed in native environments

and thus under physiological conditions (Hoerber & Miles,

2003).

Various AFM studies have been performed mostly on

biological interactions at the molecular scale and the structure

of biological membranes (Hoerber & Miles, 2003; Dufrene,

2004). In situ, short-term AFM observations of cells in their

natural aqueous environment were targeted on the structure

and functionality of cell membranes (Kamruzzahan et al.,

2004), on cell growth (Touhami et al., 2004) and division

(Matzke et al., 2001).

As previously mentioned, many environmental processes

like, e.g. cell response to environmental changes or biomineral

nucleation occur on timescales of several hours or days. Until

recently, such processes were barely investigated in situand at

nm resolution as no applicable methods were established and

tested.

Therefore it was the intention of this study to develop and

optimize an AFM approach for in vivoinvestigations on pico-

planktonic cell surfaces and their response to changes of the

chemical environment, which is an important step towards

in vivostudies on the nucleation of CaCO

3

on the cell surface.With special regard to the second step, the long-term capabilities

(> 100 h) of the approach had to be addressed and discussed

in detail as this is a prerequisite for further biomineralization

experiments in the AFM.

Cyanobacteria of the freshwater strain Synechococcus

leopoliensisPCC 7942 were chosen for the development of this

method as Synechococcus ssp. are representative of many

freshwater systems and the leopoliensisstrain is well investi-

gated from various perspectives like physiology (Wanneret al.,

1986; Arino et al., 1995; Goerl et al., 1998; Sauer et al.,

2001; Barker-Astrom et al., 2005), metabolism (Ritchie et al.,

1996; Kaplan & Reinhold, 1999; Allakhverdievet al., 2000)

and biomineral precipitation (Schultze-Lam et al., 1992;

Thompson et al., 1997). Thus, choosing S. leopoliensisPCC

7942 enabled us to compare the results with those of previous

studies by our and other groups.

METHODS

The methods, culturing and experimental conditions were

chosen with respect to the following concerns.

The first is the relevance of the experiments in order to

simulate natural processes. Second, the viability of the immo-

bilized cells had to be ensured because a proper immobiliza-

tion is necessary in order to obtain high-quality images of the

cells surface. Finally, the experimental setup has to ensure

long-term stability, which is necessary in order to attain quality

results at the designated timescale.

Cell cultivation

Synechococcus leopoliensiscells were cultured continuously in a

2.5 L chemostat with Z/10 culture medium (5.9 mg L

1

Ca(NO

3

)

2

4H

2

O, 46.7 mg L

1

NaNO

3

, 4.1 mg L

1

K

2

HPO

4

3H

2

O, 2.5 mg L

1

MgSO

4

7H

2

O, 168 mg L

1

NaHCO

3

,

11.45 mg L

1

Na-EDTA, 3 mg L

1

FeSO

4

7H

2

O, H

3

BO

3

248

g L

1

, MnSO

4

H

2

O 135

g L

1

(NH

4

)

6

Mo

7

O

24

4H

2

O

7.2

g L

1

, ZnSO

4

7H

2

O 23.2

g L

1

, Co(NO

3

)

2

6H

2

O

12

g L

1

, CuSO

4

5H

2

O 10.4

g L

1

without vitamin

solution) at a flow rate of 200 mL d

1

. The culture medium

Z/10 was chosen in order to simulate low nutrient conditions

as observed in oligotrophic lakes. Therefore, the cell physiology

of the cyanobacterial should be adapted to low nutrient

conditions. The bioreactors were aerated with sterile filtered

air. The pH of the cell suspension was measured with long-

term stable combination electrodes (InPro 3030; Mettler-

Toledo GmBH, Greifensee, Switzerland). In order to optimize

growth conditions, CO

2

was added to the aeration air when

the pH rose above the maximum value of 7.5. The cultures were

tempered to 27 C. Fluorescent lights provided a permanent

light intensity of 7 E m

2

s

1

. Cells were harvested on the

same day as the experiment was started. The cell suspension was

vacuum filtered with 0.45 m membrane filters (Schleicher &

Schuell OE67), resuspended in nanopure water and centrifugedat 2500gfor 15 min. The supernatant was drawn out of the

centrifuge tube, and the cells were resuspended in fresh

nanopure deionized water. This washing step was repeated three

times in total until the cells were used for further experiments.

Sample preparation for AFM experiments

For sample preparation, aseptic techniques were followed. All

preparation steps until mounting the sample onto the AFM


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Living under an atomic force microscope 181

2005 The Authors


scanner were performed in a clean bench (Gelair HF 36).

Round glass cover slips with a diameter of 13 mm were

precleaned with pure ethanol. After drying, the cover slips

were treated with 0.05% poly-l-lysine (Sigma Aldrich, no.

P1524) in nanopure deionized water for 48 h. The cover

slips were removed from the solution and air-dried overnight,

standing vertically on a paper towel. The cover slips were

mounted onto metal plates with adhesive carbon stickers andstored in the dark at 4 C until usage. For the cell cultivation

and cell division experiments, a glass cover slip was treated with

a drop of the cell suspension for 60 min. Then the cover slip was

vigorously rinsed with nanopure deionized water in order to

remove all cells that were not well immobilized by the poly-l-lysine.

The wet sample was mounted on the scanner of the AFM.

Gelatin-coated glass slips were prepared similar to method

described above. We followed the protocol of Doktycz et al.

(2003) using the Sigma Gelatin G6144 that gave the best

results in their study.

AFM experiments

All AFM experiments were performed in a Digital Instruments

MultiMode III AFM with a Multimode IIIa controller with

the Tapping Mode extender. Olympus Bio-Levers (type B

levers, 100 m, 0.006 N m1, sidewall angle 45) mounted

into a tapping mode fluid cell were used for the experiments.

The volume of the fluid cell is approximately 100150 L,

depending on the adjustment of the O-ring. During optimization

of the setup, Veeco NP tips (spring constants of 0.06 and

0.12 N m1, sidewall angle 35) and Ultrasharp CSC12 tips

(spring constants 0.03 and 0.05 N m1 cone angle 30)

were also tested. In comparison, the Olympus Biolevers

gave the best results because the least amount of cells was

removed during scanning. During the experiments, all scans

were performed at an angle of 90.

The AFM fluid cell was cleaned before mounting the tip and

the Z-shaped silicon O-ring, using pure ethanol. Before and

after the experiment, the O-ring was cleaned with pure etha-

nol as well and stored in nanopure water when it was not in

use. All AFM tips were exposed to intense UV light for 20 min

in order to clean the tips from potential organic contamina-

tions and to avoid bacterial contamination. All tips were used

for one experiment only.

The microscope was operated in tapping mode. The reso-

nance frequencies of the cantilevers were between 7 and9.5 kHz. Tip velocities smaller than 12 m s1 were used.

Prior to the installation of the fluid cell, the cantilever was

covered with nanopure deionized water. Z-shaped O-rings

were used to keep the cell watertight. The fluid cell of the AFM

was carefully mounted onto the sample in order to avoid

air bubbles within the cell. One of the outlets of the cell was

connected to a reservoir bottle and the other to a syringe

pump (Braun Perfusor IV, adapted to suction instead of

pressure) via previously autoclaved silicon tubes.

Cultivation experiments in the AFM

BG11 (1.491 g L1 NaNO3, 100.5 mg L1 KNO3,

35.8 mg L1 CaCl22H2O, 40.7 mg L1 Na2(EDTA)

2H2O, 74.7 mg L1 MgSO47H2O, 70.6 mg L

1 Na2HPO4,

254.3 g L1 FeSO47H2O, 152.7 g L1 ZnCl2, 1.3 mg L

1

MnCl2, 55.6 g L1 CuSO5H2O, 33.3 g L

1 CoSO47H2O,

271 g L1 NaMoO42H2O, 1.99 mg L1 H3BO3) or Z/10culture medium (described previously) was drawn through

the fluid cell at flow rates of 0.6 or 3.0 mL h1. The first

images were taken after an equilibration time of about 1 h

after mounting the fluid cell. The flow-through experiments

were run up to 6 days. In order to test the viability of the

artificially immobilized cells by an independent method, one

of the cell-treated cover slips was deposed in a sterile 50 mL

centrifuge tube with 15 mL of Z/10 culture medium and

incubated at 27 C under permanent illumination.

Cell division and surface alteration experiments

The samples for the cell division experiment were prepared

in the same way as described above. During long-term

experiments, these samples were enclosed in the fluid cell of

a Digital Instruments Multimode AFM. Several cells were

monitored in tapping mode while a solution of CaCl2/NaHCO3(with no additional nutrient concentration) was pumped

through the fluid cell by a syringe pump. The concentration of

NaHCO3 was 1.5 mM for both experiments. The concentration

of CaCl2 was 5.9 mM in the experiment where we observed the

cell division and 11.8 mM in the CaCO3 nucleation experiment.

The saturation with respect to calcite was 9.2 times for the first

case and 13.7 times for the second case. The saturation states

were calculated using PHREEQCINTERACTIVE (version 2.8) using

the wateq4f database and assuming a CO2 concentration of

370 p.p.m. in the ambient air.

Investigation of EPS by AFM in air

For investigations of the production of EPS byS. leopoliensis

PCC 7942 samples were prepared for tapping mode AFM in

air. In order to excite the production of extracellular sheath

material, samples previously used for cell division experiments

were slowly exposed to drought conditions, causing stress

on the cells. The specimen was placed in a closed Petri dish

together with a drop of water and left for air-drying for 14 days.The same microscope as described above was used for scanning.

The AFM was run in tapping mode in air with Veeco RTESP

silicon tips (125 m, 40 N m1) at a driving frequency of

318 kHz.

Sample investigation by epifluorescence microscopy

The initial samples for epifluorescence microscopy were

prepared and analysed immediately after treating the


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182 M. OBST AND M. DITTRICH

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poly-l-lysine covered glass cover slip with the cell suspension.

More samples were prepared after 1, 2 and 7 days of the flow-

through experiment (Z/10 culture medium) within the fluid

cell of the atomic force microscope.

Water was carefully removed from the glass slides using a

syringe needle. The samples were dried in an air stream. After

about 1 min, the slides were covered with a drop of immersion

oil (Zeiss 518N) and covered by a second glass cover slip. Thesamples were analysed immediately after preparation by a Zeiss

Axioscope 2 epifluorescence microscope. The microscope was

equipped with a HBO 100 W/2 mercury arc light source and

a SenSys CCD camera (KAF 1401E, 1317 1035 pixels).

CCD images were taken with a Plan-Neofluar 100. Overview

images were obtained with a Plan-Neofluar 20 in order to

characterize the homogeneity of the cell distribution. For

imaging the cell distribution and testing the viability of the

immobilized cells, the HQ filter set Cy3 (Chroma Technology

Corp.) was used. All slides were also checked for contamina-

tion (e.g. by heterotrophic bacteria) using the Zeiss filter sets

02 and 09.

TEM imaging and sample preparation

Cell suspensions for TEM experiments were prepared as

described in the section on cell cultivation. These cell suspensions

were analysed comparing three different preparation methods:

First, cell culture samples were prepared immediately after the

washing step. Next, the cells were incubated for 24 h in

1.5 mM NaHCO3 solution. Finally, the cells were incubated in

1.5 mM NaHCO3/11.6 mM CaCl2 solution for 18 and 45 h.

The final solution was 13.4 times saturated with respect to

calcite when assuming a CO2

level of 370 p.p.m. in the

ambient air. The saturation was calculated with PHREEQC

INTERACTIVE (version 2.8), using the wateq4f database. The

suspensions were centrifuged at 2000g for 10 min, the

supernatant solution was decanted. The concentrated cell

suspension was then sucked into cellulose capillary tubes,

high-pressure frozen and freeze substituted and embedded in

Epon as described by Hohenberg et al. (1994). Ultrathin

sections were prepared on a Reichert Jung Ultracut microtome

and put onto collodium/carbon-coated 300 mesh copper

grids. The sections were stained with uranyl acetate and lead

citrate according to the method cited above. Images were

taken on a Zeiss 912 enegery filter TEM.

RESULTS

Cell immobilization

In order to perform in situAFM experiments effectively, a

reproducible and reliable method for cell immobilization had

to be developed. We tested two different approaches for their

suitability for long-term experiments: immobilization on

gelatin-coated mica slides and immobilization on poly-l-lysine

coated mica and glass surfaces. In our study, the gelatin

approach did not result in a suitable long-term immobilization.

Synechococcus leopoliensiscells could not be attached to the

surface firmly enough in order to be able to perform one or

even several scans of the same area without removing the

majority of cells in this area. Furthermore, the gelatin-coated

surface often caused line artefacts, probably caused by the tip

sticking to the gelatin surface. Thus, we were not able toobtain high quality images working with gelatin-coated slips

in situ, and no images from that method are presented in this

study. For this reason, the immobilization by poly-l-lysine was

found to be superior to the former described method. When

screening several concentrations between 0.01% and 0.05%

and different exposure times, best results were obtained when

the glass cover slips were placed into a 0.05% solution of

poly-l-lysine for 48 h.

After treating the glass cover slips with a drop of the cell

suspension, the homogeneity of the cell distribution was inves-

tigated by epifluorescence microscopy. When the poly-l-lysine

coated glass cover slip was treated with freshly preparedsuspension ofS. leopoliensis, this technique resulted in a more

homogeneous distribution of cells, whereas a day old sus-

pension resulted mainly in agglomerates of cells. Thus, for each

of the following experiments, the suspension was prepared

immediately before observation, in order to obtain homoge-

neously distributed cells on the cover slips.

Cell cultivation in the fluid cell of an AFM

In order to test the vitality of the immobilized cyanobacteria,

cultivation experiments were performed in the fluid cell of the

AFM. Results of these experiments are shown in Fig. 1. In the

AFM experiments (first and second columns), the same area is

shown at each time step, whereas the epifluorescence series

(third column) shows different samples because the sampling

is disruptive. Relatively large areas of 60 60 m were

scanned several times within 6 days of cyanobacterial growth.

Because of the high tip velocity, weakly attached cells were

removed from the observed areas (compare AFM experiment

2 after 3 and after 20 h). In the AFM experiment 1 of Fig. 1,

the growth of cell clusters from a previously much lower

number of cells can be clearly observed (indicated by the

arrows). Clusters of cells could not be scanned accurately,

probably because of the production of EPS as discussed later

on. After 20 h, the first cell clusters were formed, and after3 days, the topography of the cell-covered glass slide became

too high for the z-range of the scanner and the whole image

occurred blurry. In the epifluorescence microscope the formation

of a biofilm containing several cell layers was observed. These

results totally coincide with the AFM results and explain the

difficulties of AFM imaging after several days of cell growth.

All cover slips used for the cultivation were checked for

contamination by other bacteria using different filter sets. No

cells were observed which exhibited different fluorescence


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Fig. 1 AFM cell cultivation experiments (series AFM1 and AFM2) and epifluorescence images (series epi). Each image shows a size of 60 60 m2. Series AFM1

images show the same area after starting the flow-through cultivation experiments. The arrows highlight two areas where the formation of agglomerates of freshly

divided cells (smaller size) can be observed. These cells are remarkably smaller in diameter than the ones visible at the beginning. Series AFM2 images show results

of another experiment. The formation of agglomerates can be observed until finally the vertical range of the scanner is not sufficient for imaging the area. Because

of the high tip velocity weakly attached cells were removed from the observed areas (compare AFM2 at 3 and 20 h). Series epi shows epifluorescence images of

different areas of equally treated samples. The formation of agglomerates is clearly visible until a biofilm of several cell-layers developed (138 h).


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behaviour from that ofS. leopoliensis. Thus, no indication for

contamination by, e.g. heterotrophic bacteria was found

during all long-term cultivation experiments.

A typical 3D projection of immobilized S. leopoliensiscells

during the first hours of a flow-through experiment with

culture medium is presented in Fig. 2. From the overview

scan, it is obvious that the cells are firmly attached to the

glass substrate: Almost no cells were removed during the scan

with higher magnification, which was taken first. Cells at increas-

ing progress states of cell division and separation are highlighted

by the arrows 25.

Long-term stability of scanning conditions

In order to be able to per form meaningful experiments on cell

surface alteration, in this series of experiments, on the one

hand, we had to pay special attention to the observation of

cells with respect to their vitality. On the other hand, it was

important to establish stable scanning conditions for long

periods of time. Using a solution of NaHCO3/CaCl2 without

additional nutrients facilitated these long-term experiments

as no perturbing amounts of EPS were produced by the cells

in this solution. This enabled us to find stable settings for the

feedback loop while working with the same tip for several days

(> 100 h).

Cell viability

The viability of the cells did not seem to be affected by our

technique. As shown in Fig. 2, cells at all stages of their life

cycle were immobilized on our samples. During one experiment

of 4 days, we were able to observe the continuation of a cells

life cycle. We observed the sequence of two cell divisions of an

initially rod-shaped cell into four daughter cells (Fig. 3AD).

The cell division in lateral direction was already observed when

the cell was scanned the first time, 13 h after the flow-through

experiment was started. Thirty hours after the beginning of

the experiment, we observed a line structure in the phase lag,

dividing the cell in the longitudinal axis, although at this stage,

the topographical image did not show any indication of

division. The phase lag in tapping mode is influenced mainly by

the stiffness of the sample and by adhesion forces between

sample and tip (Raghavan et al., 2000; James et al., 2001).

The reduction of the phase lag in this region therefore might

indicate an increase of stiffness in this region, most likely due

to the formation of the new cell membrane between the

evolving cells.After a time lag of about 24 h, this structure also appeared

as a constriction in the topography of the cell. This division

was then followed by little longitudinal growth of the resulting

cells. We infer that imaging the phase lag facilitates determin-

ing the onset of cell division before it becomes visible in the

topography. Figure 4 shows a 3D projection of the result of

the cell division after 101 h. The whole sequence of a cell divi-

sion was observed once. However divided, but not yet separ-

ated cells were observed more often during our experiments,

Fig. 2 AFM images showing 3D projections of Synechococcus leopoliensis

cells at the beginning of a flow-through cultivation experiment. The image of

the enlarged section (20 20 m2) was taken 2 h after the flow-through of Z/

10 culture medium was started. The overview image (60 60 m2) was taken

1 h later. Cells at different states of cell division were observed in the enlarged

image: Arrow 1 is indicating a cell division resulting in almost spherical cells. The

arrows 25 are indicating increasing progress in cell division and separation.

Fig. 3 AFM images showing a 3D projection (first column) and the phase lag

(second column) of a dividing Synechococcus leopoliensiscell at different time

intervals. Thirteen hours after the experiment was started (A), only the division

in the lateral axis is visible. Thirty hours after the beginning (B), a line structure

appears, but only in the phase-image. Twenty-four hours later (C), a furrow can

be observed in the 3D projection as well. One hundred and one hours after

starting the experiments (D) the cell division is followed by growth of the

resulting 4 cells.


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2005 The Authors


for example, during the cultivation experiment presented in

Fig. 2. An observation that is similar to the starting point of

the previously described sequence a division resulting in two

almost spherical cells ofS. leopoliensis is highlighted by arrow

1 in Fig. 2. Another example of almost spherical daughter cellsis indicated in the phase image of Fig. 7, also indicated by an

arrow.

When we analysed ultrathin sections of freeze substituted

S. leopoliensis cell suspensions from both culture broths

(Fig. 5B) and precipitation experiments in NaHCO3/CaCl2solutions and blanks in NaHCO3 solution (Fig. 5A) by

TEM, we also observed several stages of cell divisions. Most of

them resulted in rod-shaped daughter cells and some in almost

spherical cells. All divisions of elongated cells observed by

TEM were perpendicular to the longitudinal axis.

Surface alteration ofSynechococcus leopoliensis

When CaCl2 was added to the surrounding NaHCO3 solution

during our CaCO3 nucleation experiment, the surface structure

and properties of the cyanobacteria changed. The results of

these experiments are presented in Figs 3 and 6. In the 3D

projection on the left side of Fig. 6(A), the cell surface appears

rather smooth 2 h after CaCl2 was added. Only some little

protuberances can be observed. The phase image on the right

also presents a rather homogeneous surface. The line structures

parallel to the cells shape are artefacts caused by a non-ideal

shape of the tip sidewalls. Six hours after the addition

(Fig. 6B), these protuberances are more pronounced in both

the topographical image and the phase image. Twenty-onehours after CaCl2 was added to the solution (Fig. 6C), the

surface topography appears more textured than at the beginning

of the experiment. Also the phase lag is not homogeneous over

cell surface anymore.

In principle, the same observations but for different time

intervals are presented in Fig. 3. In the first image (Fig. 3A),

which was taken 13 h after the addition of CaCl2 to the solu-

tion, the surface of the cells already looks rough. Comparing

the two lower cells at the time steps of 54 h (Fig. 3C) and

101 h (Fig. 3D), the enlargement of some protuberances is

conspicuous. At this time nuclei of CaCO3 crystals on could

not be observed on the surface of the analysed cell.

Formation of extracellular polymeric substances (EPS)

The formation of a biofilm after several days of the cell

cultivation indicates that the Synechococcuscells are likely tohave started producing extracellular polymeric substances.

As EPS or sheath production is known to be strongly

influenced by the culture conditions (Merz & Zankl, 1993)

and, in particular, enhanced by drought (Potts, 1994), this

suggestion could be proven by exposure of drought stress to

the immobilized cells. In Fig. 7(A), a 3D projection of the

sample is shown. A bulky structure is visible and some

cell-sized objects seem to be embedded in this blurry matrix,

but no single cells are visible. In contrast to the height data,

the cells are clearly visible in the phase image of this sample

(Fig. 7B). The suggestion of cells embedded in a matrix is

confirmed by this result. In air, the EPS becomes rigid, enoughto form a surface that can be scanned by AFM. However, it was

very difficult to scan samples covered by EPS in an aqueous

environment. Figure 8 shows a topographical image of the

same area during a flow-through experiment at different time

intervals. After 2 h of pumping culture medium through the

fluid cell, a good image quality was achieved (Fig. 8A), whereas

after 4 h (Fig. 8B), the first artefacts were observed. However,

17 (Fig. 8C) and 18 h (Fig. 8D) after beginning to culture

cyanobacteria within the fluid cell of the microscope, no

parameter set for the feedback loop could be found in order

to run the microscope under stable conditions. The images

contained many line artefacts, probably caused by the tip

occasionally sticking to a bulk of EPS.

DISCUSSION

Influence of culturing and experimental conditions

on cyanobacteria

With respect to both the design of the long-term in vivo

experiments in the AFM and the relevance of the experiments

for simulating natural conditions, the culturing conditions

had to be considered carefully. BG11 is a widely used medium

for the cultivation of cyanobacteria. This medium, which was

originally optimized for the cultivation of coccoid cyanobacteria(Stanier et al., 1971), is rich in both nitrate and phosphate.

It is not representing conditions that are observed in oligotrophic

freshwater systems, which are often phosphate-limited. For

some sensitive phycoerythrin-rich red pigmented Synechococcus

species, BG11 was even shown to slow down growth compared

to lower concentrated media (Ernst et al., 2005). Thus, culturing

and experimental conditions are influencing physiology,

ultrastructure and metabolism of the cells (reviewed by

Allen, 1984; de Marsac & Houmard, 1993); therefore surface

Fig. 4 AFM 3D projection of the divided cells after 101 h. The fourfold division,

which is caused by two subsequent divisions perpendicular to each other, is

clearly visible.


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2005 The Authors


properties of the cells as well (Borroket al., 2004; Haas, 2004).

When the experimental conditions were not representative of

those in nature, the relevance of the experiments with respect

to simulating and understanding natural processes would be

strongly affected. One example with respect to our experiments

is the production of EPS by the cells. Both composition and

production were shown to be strongly dependent on: (i) the

physiological state and growth state of the cells, (ii) the

composition of the nutrient media, and (iii) the ionic and

physical conditions of the media (reviewed by Decho, 1990).

Another more ecological aspect is the importance of pico-

plankton within the phytoplankton community. Picoplankton

can dominate the primary production especially in oligo-

trophic lakes (Stockner et al., 2000). Nutrient concentrations

Fig. 5 TEM images showing the ultrastructure of variously treated Synechococcus leopoliensisPCC7942. Cell viability was not affected after 24 h of treatment in

1.5 mM NaHCO3 solution without addition of other nutrients (A), as cell divisions were observed in this solution. However, starting separations of the thylakoids

were observed (indicated by S), these are the first indications of adaptation to these conditions. An almost finished separation of two cyanobacterial cells in culture

medium Z/10 is presented in the figure (B). After 45 h in NaHCO3/CaCl2 solution (C), which is used for the experiments on surface alteration and CaCO3 nucleation,

the thylakoids of the cells are intact. However, cell metabolism adapted to the nutrient-poor conditions by accumulating glycogen granules (indicated by G).


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2005 The Authors


and temperature were shown to control the contribution

of picoplankton to phytoplankton biomass and production

(Agawin et al., 2000a,b). Therefore, the environmental

relevance of the processes we investigate in the laboratory

is strongly linked to the experimental conditions. In order to

represent these conditions and to culture a sufficient amount

of cells for our experiment in reasonable time, we decided to

culture S. leopoliensisin Z/10 medium. The major nutrients:phosphate, nitrate and sulphate are 25, 33 and 30 times less

concentrated than in BG11-based media that were used for

many experiments, e.g. studies on sulphur starvation (Arino

et al., 1995) or nitrogen starvation (Sauer et al., 2001; Barker-

Astrom et al., 2005) or both (Wanner et al., 1986). Thus, the

metabolism and therefore the growth rate of our cultured

cyanobacteria were adapted to these conditions. This is indi-

cated by a specific growth rate that is reduced by more

than one order of magnitude in continuous Z/10 cultures

compared to BG11 batch cultures. We measured equilibrium

growth rates of 0.1 d1 in our chemostats (light intensity

7 E m2 s1), compared to measured maximum -values

between 1 d1 and 2 d1 (average 1.8 d1, light intensity 12 E

m2 s1) in BG11 medium in batch experiments. However, the

nutrient concentrations were sufficient in order to maintain

photosynthetic active and viable cells. The cultures showed the

typical blue-green colour. No indications of starvation-inducedchlorosis could be found as described in Sauer et al. (2001)

and Goerl et al. (1998). In their studies, the authors describe

the acclimation and the loss of photosynthetic pigments as an

adaptation to long-term survival. Concerning growth, our

results also coincide with the findings of Timmermans et al.

(2005). They demonstrated that small phytoplankton species

can grow under much lower nutrient conditions and concluded

that picoplankton will hardly ever stop growth totally by nutri-

ent limitation under natural conditions. Collier & Grossman

Fig. 6 AFM images showing a 3D projection (first

column) and the phase lag (second column) of the

surface alteration of a Synechococcus leopoliensis

cell at several time intervals after the cell was

exposed to Ca2+: 2 h after the beginning of the

experiment (A), the surface appears rather smooth

in both, topography and phase. After 6 h (B) small

protuberances are more noticeable and after 21 h

(C) the cell surface seems to be rather rough in

topography. The protuberances are still clearly

visible in the phase image.

Fig. 7 Three-dimensional projection (A) and phase

image (B) of Synechococcus cells embedded in

huge amounts of EPS. While it is not possible to

identify the cell structures within the bulk mass of

sheath material in the topographical image, they

can easily be identified in the phase image. Theseimages were taken in tapping mode in air. The

arrow is indicating a previous cell division resulting

in almost spherical daughter cells.


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(1992) measured continued cell division ofS. leopoliensis

PCC 7942 in their nutrient deprivation experiments, but

depending if P, N or S was depleted, the time steps between

cell divisions increased to certain extents. Thus, the expression

starvation, which is used in most of the studies cited above,

can easily be misinterpreted to some extent. The expression

adaptation describes the process more accurately as supported

by the studies of Sauer et al. (2001) and Goerl et al. (1998).

Another aspect, the influence of nutrient depletion on cell

physiology, morphology and ultrastructure of cyanobacteria,

was proven carefully in our study. Ultrastructural changes

affecting the cell wall were not observed in any of our experi-

ments. Hardie et al. (1983) describe in detail several steps of

intracellular changes due to iron starvation. The first step is the

separation of the thylakoids and was observed between 16 and

60 h of iron starvation. Degradation or deterioration was not

observed in any of their cells at this stage. However, after >60 hof starvation, they observed deterioration of the thylakoid

system and in some of the cells a separation of the cytoplasmic

membrane and the peptidoglycan layer. This separation could

change the morphology of the cells and was never observed

during the first step. Iron starvation did not affect carboxysomes

or the extracellular glyocalyx in their experiments.

The reversible disintegration of the thylakoids membranes

was also observed after 9 days of sulphur starvation experiments

on Gloeothecesp. PCC6909 by Arino et al. (1995).

In contrast to their results, these ultrastructural changes

were observed to much less extent during our experiments.

In Fig. 5(A) (taken after 24 h in NaHCO3 solution), the thy-

lakoids are less separated than in Fig. 2(A) of Hardie et al.

(1983), which is described as an early stage of separation. For

our experiments, Fig. 5(A) is representative as we observed

this stage for the majority of the cells we analysed by TEM

(>15).

Furthermore, we found that Ca2+ most probably plays an

important role in the stability of the thylakoid system as the

separation was quite reduced when S. leopoliensis cells were

suspended in NaHCO3/CaCl2 solution even after extended

periods of time (45 h in Fig. 5C) compared to the Ca-free

solution of NaHCO3. In the Ca-containing solution, the vast

majority of the cells did not indicate separations of the thyla-

koids (>15 cells analysed). Thus, morphological changes of

the cell and, therefore, of the cell surface due to starvation arevery unlikely to happen in the timescale and under the condi-

tions of our in vivoAFM experiments. Only the very first steps

of this process were observed by TEM in an even more nutri-

ent depleted solution than the one used for the nucleation

experiments.

A noticeable ultrastructural change that occurred in the

NaHCO3/CaCl2 sample compared to the cell from the cul-

ture medium is an accumulation of glycogen granules that

makes the cell appear more electron translucent. Our findings

Fig. 8 The production of sheath material during

an in situ flow-through AFM experiment is shown

for several time intervals. Two hours after the

cultivation experiment was started (A) the image

is rather clear, many cells of small diameter wereobserved in the middle of this image. Six hours after

starting (B) the cell agglomerate can still be easily

identified. Twenty-one hours after the start (C) the

image was already blurry as the tip started sticking

to the sample. Twenty-two hours after starting

cultivation (D), it is not possible any more to

identify single cells within the blurry mass of sticky

sheath material.


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2005 The Authors


coincide with the results of Wanner et al. (1986). In Fig. 2 of

their study, they present very similar cells of the S. leopoliensis

PCC 6301 strain, which were observed in stagnating and in

nitrate-starved cultures after 9 h of starvation. Size and shape

of their cells did not change significantly even after extended

periods of starvation (70 h). These results coincide with our

findings.

To sum up, the results of this study emphasize the impor-tance of the culturing and experimental conditions. As this was

done extensively in the case of our study, we could show the

relevance of our first results with respect to simulating natural

processes in order to clarify the mechanisms. The reason why

we cannot find strong indications for starvation even after 45 h

in nutrient-free NaHCO3/CaCl2 solution, is most probably cell

metabolisms, which are already adapted to low nutrient

conditions (and slowed down compared to BG11 cultures)

because of the cultivation conditions in Z/10 medium.

Cell immobilization

The limiting factor of cell imaging is a proper immobilization

method that attaches the cells firmly to the substrate but avoids

denaturizing the sample (Dufrene, 2004). Some microscopists

tried to work around this problem when they performed

experiments in situand then dried the samples for the AFM

imaging. Camesano et al. (2000) for example covalently bound

bacteria to the support. They could perform experiments with

the immobilized cells in an aqueous environment and air-dry

the samples for imaging. Liu et al. (2004) also performed

AFM experiments in air and observed structural differences

in the cell membrane between La3+ treated Escherichia coli

cells and nontreated ones. Gently air-dried polysaccharide

macromolecules are suggested to keep their hydration water

(Santschi et al., 1998) but it is not clear if biopolymers maintain

their conformation (Wilkinson et al., 1999). Bolshakova et al.

(2001) reported two kinds of artefacts induced by drying

biological samples when they compared images of bacteria

scanned in air and in situ, respectively. First, the height and

width of the cells decreased when dried. Second, a surface

pattern appeared in air, which could not be observed when

imaging in water. Therefore, cell imaging in water seems to be

superior with respect to an easier interpretation of the obtained

data.

However, scanning cells in an aqueous environment is

still a challenge (Dufrene, 2003) because of the delicate im-mobilization. The optimization of the immobilization is

necessary as an improvement of cell adhesion was indicated to

promote cell resistance to the disruptive effect of the scanning

cantilever (You et al., 2000) and, therefore, reduces artefacts.

In principle, five different immobilization methods have been

established. (1) Mechanical trapping of cells in the filter pores,

e.g. an isopore polycarbonate membrane (Touhami et al.,

2003, 2004). (2) Physical adsorption of cells onto chemically

treated surfaces, e.g. poly-l-lysine (Bolshakova et al., 2001;

Doktycz et al., 2003) or polyethylenimine-coated supports

(Crawford et al., 2001). (3) Embedding cells into a soft

layer of agarose (Gad & Ikai, 1995) or gelatin (Doktycz et al.,

2003). (4) Direct growing of the cells onto a suitable sub-

strate (Kuznetsovet al., 1997; Gebeshuber et al., 2003). (5)

Drying of cells on a surface of an aluminium oxide anodisc

filter for several hours and the following rehydration of the

cells (Yao et al., 2002).All of these methods have advantages and disadvantages

that will be discussed in the following section with special

regards to their potential of performing long-term in vivo

experiments. Vadillo-Rodriguez et al. (2004) compared the

first two methods and concluded mechanically trapping

bacterial cells in filter pores to be more reliable. This method

has at least three possible drawbacks. First, it is not applicable

to rod-shaped micro-organisms. Second, EPS might accumulate

on top of the anchored cells, and third, the rigid entrapment

of cells in holes of limited diameter might affect the viability

of the cells (Gad & Ikai, 1995). Thus this method might be a

good option for short-term in vivo investigations of un-disturbed cell surfaces. Doktycz et al. (2003) compared the

immobilization of different bacterial strains by adsorption

onto different poly-l-lysine coated surfaces to different

gelatine-coated ones. In their study, gelatin turned out to be

superior to poly-l-lysine as it attached the cells more firmly.

Both methods are capable of immobilizing rod-shaped cells.

The cell surface at the interface between cell and substrate

might be artificially altered by linker molecules like poly-

l-lysine. However, as the thickness of the coating of the

substrate is very little (on the molecular level in case of

poly-l-lysine), the AFM-accessible parts of the cell are likely

to be unaffected.

The fourth method is restricted to organisms or cells, which

tend to grow on solid substrates, but it is not applicable for

example on single-cellular organisms living in a water body.

The last of the five techniques was described to retain a suf-

ficient number of Gram-negative bacterial cells attached to

the membrane surface in order to perform experiments in an

aqueous milieu for hours. The authors claim that this method

was developed particularly considering the minimization of

surface alterations, which might be caused by binding agents.

However, drying of the cells might cause changes of the cell

metabolism and enzyme activity (e.g. Potts et al., 1984) and

surface alterations due to membrane modifications in particu-

lar. A review of observed cell responses to desiccation can befound in Potts (1994). An alteration of the surface, however,

would be critical with respect to the surface-related nucleation

processes that we wanted to observe in our experiments.

Beside these alterations, another cell response is limiting this

approach: In his review, Potts (1994) reports that many

cyanobacterial strains secrete conspicuous amounts of EPS

when they are exposed to water stress. As discussed later in

this chapter, EPS is one of the major limiting factors when

performing long-term experiments in the AFM.


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With respect to our requirements, we proved that the

second method using poly-l-lysine was superior to the third

method using gelatin for our experiments. Poly-l-lysine-coated

glass cover slips were shown to be suitable for immobilizing

single-cellular strains of cyanobacteria for in situAFM investi-

gations in tapping mode. In contrast to the findings of

Doktycz et al. (2003), who investigated the immobilization

ofStaphylococcus aureus (Gram-positive) and E. coli (Gram-negative), in our study with S. leopoliensis(PCC7942), poly-

l-lysine was found to be superior to gelatin. Using gelatin, we

could immobilize cells only for short periods, but after one or

a few scans, the majority of the cells was shifted away from

the tip. Another disadvantage of the gelatine method was the

frequent occurrence of line artefacts in the background. These

artefacts were most probably caused by the tip sticking in the

soft gelatin layer that covered the glass substrate. Thus, it was

rather difficult to find the right settings in order to establish

a stable feedback loop. With gelatin, high-quality images

of S. leopoliensis could not be recorded. In contrast to this

method, the poly-l-lysine-coated cover slips were shown to becapable of immobilizing S. leopoliensiscells properly for peri-

ods longer than 100 h. During this period of time, the cell

shown in Fig. 3 could be scanned more than 20 times in total!

The preparation mode and age of the used cell suspension

have an influence on the homogeneity of the cell distribution.

The preparation can be optimized using epifluorescence

microscopy as an effective tool for analysing the distribution of

cells on the cover slips. ImmobilizedSynechococcuscells rapidly

exhibited cell division and growth without a pronounced

lag phase after starting the experiments in growth medium.

This result provides additional evidence that cyanobacteria

cultivated in homogeneous suspension are viable under

biofilm conditions (Becker et al., 2004). Thus, we conclude

that immobilization of cells on flat surfaces by poly-l-lysine is

suitable for long-term experiments. Especially for in situinves-

tigations of the cell surface under changing environmental

conditions, this method is superior to physical entrapment and

embedding in agarose or gelatin. However, this method is

most probably restricted to cells with negative surface charges

and most likely has to be optimized for each individual strain.

First results of long-term surface alteration experiments

After culturing conditions and cell immobilization were

optimized, we could successfully perform initial long-term invivoexperiments on the changes of cyanobacterial surfaces due

to the treatment with supersaturated solution with respect to

calcite. The relevance of these results for natural processes was

ensured as discussed in detail in the preceding section.

Ca2+ ions were observed to alter the surface of the cells

significantly within the timescale of hours in a NaHCO3/CaCl2solution. These alterations do not only change the surface

microtopography ofS. leopoliensisbut also the physical prop-

erties of the cell surface as the changes could be observed in

the phase lag as well. Nuclei of initial CaCO3 crystals could not

be identified on the analysed cell. However, from SEM analysis

of precipitates of bulk experiments (data not shown), we know

that only a fraction ofS. leopoliensiscell nucleates calcite on its

surface. Thus, we conclude that a series of experiments has to

be performed until the sequence of crystal nucleation can be

observed.

The observed changes in the fine structure of the cyanobac-terial cell surface, however, could be shown not to be caused

by starvation, but by Ca2+ ions of the supersaturated solution.

This is supported by the following results: The cell surface

observed by AFM is rather smooth in Z/10 culture medium

(Fig. 2), pure NaHCO3 solution (data not shown here) and

at the beginning of the nucleation experiments after 2 h in

NaHCO3/CaCl2 solution (Fig. 6A). The most advanced steps

of the starvation process in our experiments, however, were

observed by TEM after treatment in pure NaHCO3 solution.

These changes were described as early stages of the separation

of the thylakoids in the work of Hardie et al. (1983). This

stage is described as the initial step of an iron starvationprocess and still far away from affecting the structure of the

cell wall.

However, the physicochemical principles of the observed

surface changes during our nucleation experiments could not

be clarified in this first study. More advanced techniques like

chemical force microscopy could help in order to characterize

the changes more in detail.

Cell division

Another but rather unexpected result was the observed second

cell divisions perpendicular to the first divided cells (Fig. 3),

as Synechococcususually divides perpendicular to the length axis

as shown in the TEM image of Fig 5(A,B). The starting points

of the unusual division are two almost spherical cells that seem

to be already divided, but not yet separated. The unusual divisions,

perpendicular to the first one, to the authors knowledge have

never been observed before for S. leopoliensis. This observation

was only made once during our long-term nucleation experiments.

However, it is rather unlikely to observe ongoing cell division

online, simply based on statistics. In Z/10 medium, the

specific growth rate of our culture was 0.1 d1 at 27 C and

an illumination of 7 E m2 s1. With regard to the study of

Collier & Grossman (1992), we suppose the growth rate to

be smaller in NaHCO3/CaCl2 solution. Nutrient supply islimited at one side of the cell because of immobilization and,

furthermore, the temperature was only 20 C. The illumination

in the fluid cell cannot be measured with conventional

instrumentation. Additionally, the lateral resolution needed

to be sufficient in order to be able to recognize cell divisions,

i.e. the scan area has to be small, the resolution settings high

(512 512 pixels) and the scan speed slow. Combining these

premises, it is evident that the probability of observing ongoing

cell divisions is rather small. Completed cell divisions of


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2005 The Authors


rod-shaped cells were observed frequently as shown for

example in Fig. 2. Although, divisions resulting in spherical

cells were only observed a few times during our experiments,

they were observed for both immobilized cells by AFM and

suspended cells in the free water by TEM.

However, reasons other than cell divisions can barely explain

our observations. Contamination by other strains is not likely

as aseptic techniques were followed as far as possible, a pureculture was used for these experiments and contamination

was never observed when analysing the samples under the epi-

fluorescence microscope. The simple loss of cell pressure and

collapse of the cell for reasons of starvation were, on the one

hand, shown to be unlikely and, on the other hand, this proc-

ess would hardly lead to the observed fourfold structure in

topography (Fig. 4). It has been shown by Begg & Donachie

(1998) that round sister cells of an usually rod-shaped E. coli

strain divide at alternating planes at 90 to each other, when the

physical separation of the sister cells is prevented by immobi-

lization on agar. Therefore, we strongly believe that the observed

changes of our S. leopoliensiscell represent two subsequentdivisions of a single cell along perpendicular division planes.

Possible artefacts

When soft and fragile samples like cells are scanned by AFM,

possible artefacts induced by multiple scans of the same area

have to be addressed. Especially when we scanned large areas

for overview images at high tip velocities (e.g. 60 60 m2 in

Fig. 1), some cells were removed from the surface simply due

to lateral forces applied by the scanning tip. Scanning smaller

areas allowed a significant reduction of the tip velocity and

still a reasonable acquisition time. Thus, scanning small areas

was found to be more convenient when working in aqueous

environments. Working at a scan-angle of 90 also improved

the image quality, because adherence artefacts only lead to

torsion of the cantilever, but influences the feedback-loop of

the AFM to a lesser extent. During the optimization of our

setup we also observed that using tapping mode facilitates the

long-term immobilization of our S. leopoliensiscells. Whereas

in contact mode the cells were frequently removed by the tip,

this problem was greatly reduced when working in tapping

mode. In comparison to contact mode, tapping mode applies

less shear stress to the sample. The shear stress in contact

mode was shown to cause artefacts as disruptive effects of the

cantilever in the study of You et al. (2000). Finally, artefactsdue to the tip geometry have to be considered when relatively

large objects in height-like cells are scanned. The upper surface

of the cells can be interpreted without problems, but the

sidewalls are affected by the tip geometry (Velegol et al., 2003).

Long-term capabilities of the in vivo AFM approach

Concerning the long-term capabilities of our approach, several

important topics have to be addressed. Cell viability and

immobilization have already been discussed with respect to

long-term experiments in the preceding sections. Another

important topic is the stability of the scanning conditions. The

microbial production of EPS turned out to be the limiting key

factor, deciding if high-quality images of immobilized cells

could be obtained or not. In a nutrient-depleted medium like

the NaHCO3/CaCl2 solution used for our experiments, it was

possible to adjust feedback parameters of the AFM accuratelyin order to have a stable feedback loop even after several days.

In culture medium, this was not possible with our setup,

basically because of the production of loosely bound EPS by

the cyanobacteria. Whereas in situAFM is highly capable of

investigating thin mucilage layers that are strongly attached to

cells (e.g. Higgins et al., 2003), it is hardly possible to

elucidate the microtopography of thick, soft bulk exopolymers

produced by cyanobacteria (e.g. Fig. 8). The tips stuck to the

EPS frequently so that the induced line artefacts prohibit

reasonable interpretation of the obtained images. We emphasize

that experimental conditions have to be optimized not only

with respect to the cell viability and relevance for naturalprocesses, but as well as for the quality of the scanning

technique. In the case of our study, these optima luckily fit

together. However, in the case of other geo-related in vivo

AFM studies, the optimization might require compromises.

CONCLUSION

For the first time, it is possible to investigate processes at the

interface between biology and geology on a relevant timescale,

in vivoand nearly online. The possibility of performing long-

term experiments on bacterial surfaces under nearly environmental

conditions offers great opportunities for investigations on

surface-related biogeochemical processes like biomineral

nucleation or metal adsorption. We tried to cover both potentials

and possible drawbacks of this approach in our study and we

hope to contribute some reassurance for future requirements

of long-term in vivoAFM studies.

ACKNOWLEDGEMENTS

This study was ETH funded (TH 0-20967-02). We are grate-

ful to M. Boller (Eawag) for loaning the AFM. We would like

to thank B. Wehrli, B. Sinnet (Eawag), Dr S. Akari and A. Korte

(NanoCraft) for useful and critical discussion of the results.

Finally we thank the reviewers for improving the quality of thismanuscript.

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