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* Department of Physics and Astronomy, Vanderbilt University, Nashville, Tennessee 37235; Institute for Botany, Plant Biophysics, Darmstadt University of Technology, D-64287 Darmstadt, Germany; and Physics Department, Institute of Mathematics, Physics and Mechanics, University of Ljubljana, 1000 Ljubljana, Slovenia
Correspondence: Address reprint requests to Zvonko Trontelj, Physics Department, University of Ljubljana, Jadranska 19, 1000 Ljubljana, Slovenia. Tel.: 386-1-476-6582; E-mail: zvonko.trontelj{at}fiz.uni-lj.si.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSANALYSIS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSANALYSIS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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; Hope and Findlay, 1964; Hope and Walker, 1975; Tazawa et al., 1987). Experimental work was favorable given the very large geometry of a single internodal cell and the many possibilities of manipulating these cells with microsurgery (Shimmen et al., 1994; Beilby, 1990). In this sense, the species Chara corallina is the plant equivalent to the squid axon in the study of ion transport in plants.
The recent phylogenetic analysis of the Characeae has again strengthened the model character of Chara. This family of green algae is now seen as the ancestor of higher plants (Kranz et al., 1995; Karol et al., 2001). Hence, an investigation of the physiological processes in Chara is interesting from an evolutionary point of view. Also, data from the simple single-cell system can be seen as a common primitive mechanism for comparable effects in higher plants.
The plasma membrane of Chara corallina is electrically excitable. Depolarization of the membrane more positive than a critical threshold elicits a propagated action potential (AP) (Beilby and Coster, 1979). The bulk rise in membrane conductance, which underlies this transient membrane depolarization, is due to a short-lasting activation of Cl– and K+ channels (Homann and Thiel, 1994; Thiel et al., 1997). Because of the thermodynamic conditions, this leads to an efflux of KCl from the cytoplasm into the external medium (Kikuyama, 1986). Hence, the plant action potential is different from the AP in animal cells; it is osmotically active. This osmotic activity seems to be fundamental for the physiological role of the AP in Chara where it is apparently involved in the regulation of the internal pressure (turgor pressure) in the cell (Barry 1970; Shepherd et al., 2002).
The mechanism underlying the electrical stimulation of the AP is still not fully understood, but a rise in cytoplasmic Ca2+ undisputedly plays a key role (Williamson and Ashley, 1982; Kikuyama and Tazawa, 1983; Wacke and Thiel, 2001). This rise in Ca2+ is thought to activate Ca2+-sensitive Cl– channels (Okihara et al., 1991; Homann and Thiel, 1994)—the process that generates the depolarization. The activation of K+ channels, which support repolarization, follows either as a result of the depolarization or in response to the rise in Ca2+ (Homann and Thiel, 1994; Thiel et al., 1997).
The classical view is that Ca2+ enters the cytoplasm via voltage-dependent channels (Tazawa and Kikuyama, 2003). Other investigations have revealed that the threshold for excitation is posed by a quasi all-or-none type liberation of Ca2+ from internal stores (Wacke and Thiel, 2001; Wacke et al., 2003). In this sense, the AP in Chara occurs not to function like a classical Hodgkin Huxley (HH) type AP. This means that the AP is not entirely based on the time- and voltage-dependent activation properties of plasma membrane ion channels but on a complex signal transduction cascade. Similar mechanisms of membrane excitation, which are based on Ca2+ release from internal stores, are also well known from animal cells where they are found in muscles (Nelson et al., 1995) and even some neurons (Chavis et al., 1996).
The latter type of a "chemical" action potential was in the past well-described by models, which include a nonlinear dynamic interplay of cytosolic Ca2+ ([Ca2+]c) and second messenger-stimulated release of Ca2+ from internal stores (Othmer, 1997). The same modeling approach was also suitable to simulate a large spectrum of phenomena related to membrane excitation in Chara (Wacke et al., 2003). One parameter in this model, which is predicted to effect the kinetics of Ca2+ mobilization and hence the kinetics, is the cytoplasmic concentration of Ca2+ before stimulation.
To further prove the validity of the model, we therefore examine in this study the kinetics of the AP under conditions in which [Ca2+]c is altered. This can be easily done by transferring the plants from the dark into the light, because it is known that [Ca2+]c is in these cells reduced under the influence of light (Miller and Sanders, 1987; Plieth et al., 1998a).
This work also has a methodological aspect because the AP in Chara is examined with a noninvasive method by recording the magnetic field in the vicinity of a C. corallina internodal cell under different illumination conditions. According to Ampere's or the Biot-Savart law, measured magnetic field in the vicinity of intra- and extracellular currents carries information on these currents. Measurement of cellular magnetic field is also connected to the AP (Clark and Plonsey, 1966; Woosley et al., 1985) and can be thus a supplemental method to other electrophysiological investigations.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSANALYSIS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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60 cm high) filled with artificial pond water (APW) (0.1 mM KCl, 0.5 mM CaCl2, 1 mM NaCl). The plants were grown on a layer of lake mud. A single internodal cell was prepared a day or two before the experiments and placed in APW + 2 HEPES, pH = 6.8–7.0 (NaOH). All salts were from Sigma Chemical (St. Louis, MO).
The internodal cell was held in a horizontal Plexiglas half-tube, similar to that previously described (Trontelj et al., 1994) and is schematically shown in Fig. 1 (side view) and Fig. 2 A (top view) . The leveling stage allowed the bath to be moved up and down to adjust the distance between the cell and the tail of the superconducting quantum interference device (SQUID) microscope Dewar vessel. A 10 µm thick Mylar film prevented contact between the cell and the SQUID microscope sapphire window and stabilized the position of the internodal C. corallina cell during scanning. One end of the internodal cell was mounted in a small Plexiglas compartment so as to electrically isolate this end of the cell (the left end in Fig. 1) from the bath with a petroleum jelly seal (Fig. 1). The Plexiglas compartment was filled with 100 mM KCl that served as a reference electrode (the K+ anesthesia technique) for parallel electrical measurements of the AP (Trontelj et al., 1994). We had to apply this technique so as not to disturb the SQUID sensor with ferromagnetic impurities of microelectrodes and the cell manipulating device. The Plexiglas half-tube with the internodal C. corallina cell and the 100 mM KCl compartment was submerged in a shallow Plexiglas container with the dimensions 20 x 14 x 4 cm3 (Fig. 1). Using Ag/AgCl electrodes, we measured the resting potential as the potential difference between the APW and the KCl compartment before stimulation. Before the measurements, both recording electrodes were immersed in the KCl compartment to cancel the junction and offset potential. We needed C. corallina internodal cells that were at least 8 cm long to allow simultaneous measurement of the electric potential differences (AP) and the associated magnetic field due to action currents (AC). For the measurements reported here, we selected internodal cells that were up to 15 cm long with diameters between 0.8 and 1.1 mm. The amplitude of the square wave electric stimulus with duration of 40–80 ms was adjusted above the threshold for the AP eliciting. The interstimulus interval was at least 120 s, and most cells were responsive to several tens of stimulations.
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The cell illumination was provided by a light source with a spectrum similar to daylight with 5–10 Wm–2 delivered to the cell by a fiber-optic bundle that passed through an opening into the magnetically shielded room to eliminate power supply noise of the light source.
By measuring magnetic fields in the vicinity of a C. corallina internodal cell after the AP was electrically elicited, we basically measure the contribution of total ionic current, associated with the propagating AP. The measured internodal C. corallina cell is in APW bath, and extracellular ionic currents, caused by the AP propagation along the cell, are spread through the whole APW volume. Their density is very small and their contribution to the measured magnetic field is negligible. Practically only the magnetic field due to axial intracellular current—we can call it the action current—can be measured. Taking the maximal intracellular current 1 µA and the distance from the cell geometrical axis 2 mm, we obtain by applying Ampere's law for the maximal value of magnetic field 10–10 T = 100 pT. To measure this magnetic field or even smaller magnetic fields, a very sensitive magnetic field sensor is needed. We can choose between a SQUID magnetometer (Drung, 1995), covering an area of several 100 cm2 and a single SQUID magnetometer with a small detection coil (usually called a pickup coil) in a configuration that comes extremely close to the measured object—SQUID microscope (Baudenbacher et al., 2002). The second alternative enables us to obtain a high signal/noise ratio (small distance from the detection coil to the cell) and high spatial resolution (due to the small diameter of detection coil).
In our SQUID microscope design, a hand-wound niobium detection coil is coupled to a commercially available low temperature SQUID sensor. The SQUID sensor and the detection coil are in the vacuum space of the cryostat separated typically by 100 µm from the room-temperature sample by a thin sapphire window. A computerized nonmagnetic scanning stage with submicron resolution in combination with a tripod leveling system allows samples to be scanned at levels of 10 µm below the sapphire window. For a 20-turn 500 µm diameter detection coil, we achieve a field sensitivity of 350 fT/Hz–1/2 and for a 40-turn 1 mm diameter coil, 50 fT/Hz–1/2 for frequencies above 1 Hz for the vertical component of the magnetic field. The voltage output of the SQUID electronics corresponding to the vertical magnetic field component generated by propagating AC was recorded at a bandwidth of DC-500 Hz for a period of 30 s at a position along the cell where the magnetic signal was maximal. The SQUID microscope, the sample, and the scanning stage were housed in a three layer, µ-metal magnetically shielded room (Vacuumschmelze, Hanau, Germany) to eliminate the influence of near-DC and higher frequency noise sources.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSANALYSIS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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200 µm above the C. corallina internodal cell (Fig. 2 A) where the vertical component of the magnetic field was readily detectable (at least several 10 pT at maximum). The sapphire window of the SQUID microscope was positioned 100 µm above the cell surface, resulting in a distance between the detection coil and the cell surface of 200 µm. The signal/noise ratio was 5:1 or better for a single scan. A high temporal density of measurement points allowed us to improve the signal/noise ratio by averaging. We regularly confirmed that the signal reversed polarity (as expected for this geometry) after the scan crossed the geometrical axis of the cell. Keeping the SQUID magnetometer's detection coil in position, denoted as "Site 1" in Fig. 2 A, we measured the magnetic field as a function of time. The result is shown in Fig. 3 A. Moving the bath with the C. corallina internodal cell so that the detection coil reaches position "Site 2" in Fig. 2 A causes the signal polarity to change (Fig. 3 B). Small differences in peak amplitudes between Fig. 3 A and B reflect scattering between two measuring sessions; a small deviation from symmetrically located measuring points possibly contributes to these differences as well.
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4 cm/s, which is in good agreement with the previous results (Trontelj et al., 1994). Before and during these measurements, the C. corallina internodal cell was kept in darkness for at least 1 h. The detection coil was placed 2 mm (Fig. 3 B) and –2 mm (Fig. 3 A) lateral to the internodal cell surface. The vertical distance from the detection coil to the cell surface was 200 µm. The measurements shown in Fig. 3 are a single-shot measurement with no averaging of the data. The dotted line in Fig. 2 A denotes a small part of the internodal cell, which we assume is sufficiently long to cover both the repolarized and the depolarized area in this part of the internodal cell. The corresponding AC is in the form of two current dipoles (called also axial current elements) and is drawn as two thick arrows within the cell (Fig. 2 B). The left one belongs to the depolarized part and the right one to the repolarized part of the cell. The magnetic field around each current element is for clarity shown as one broad line in the ring form around the current element according to Ampere's law. Both magnetic fields have different directions since the current elements are pointing in opposite directions. Considering the constant AP propagation velocity c of 4 cm/s and the duration of depolarization 1 s, we get for the depolarized part of the cell x = ct = 4 cm. This means that for measurements very close to the C. corallina internodal cell, we have a similar situation as in the case of observing the magnetic field of a straight wire that is conducting electric current.
The relation between AP and AC (and corresponding magnetic field) in different nerve cells (Barach et al., 1980; Woosley et al., 1985; Trahms et al., 1989) and also in an excited single internodal cell of C. corallina (Slibar et al., 2000; Trontelj et al., 1994) was studied in detail. Briefly, in all studied cases, we observe that the peak forward action current occurs approximately simultaneously with the maximum time derivative of the AP, and the maximum backward action current occurs at the most rapid change in the AP during repolarization. The time of the peak AP corresponds to the zero-crossing between the forward and backward phases of the AC. The correlation of these AP and AC features are more pronounced in our measurements in this study since the small SQUID microscope detection coil can be placed closer to the cell. Fig. 3 illustrates these relations. However, we have to take into account in this particular case that AP and magnetic field were not measured at the same point along the cell (there is 1.1 s time difference between both signals).
Under constant illumination and temperature, the transmembrane potential and the magnetic field generated by AC revealed only small variations either when different cells or when one C. corallina internodal cell was measured repetitively.
Fig. 4, traces a–i, show the time dependence of the vertical component of magnetic field measured at y = 2 mm lateral to the cell's geometrical axis under different illumination conditions. In this series of experiments, the internodal cell was in darkness for 60 min before the first measurement of AP propagation. Fig. 4, trace a, shows the magnetic field generated by AC just before the light was turned on. Subsequent recordings were done under constant illumination after particular time intervals as shown in Fig. 4, traces b–e. It can be seen that, as a response to illumination, the temporal evolution of the magnetic field changes over a period of <1 h before reaching a new steady state. Most pronounced is the time shift of the positive peak of the magnetic signal associated with the depolarization. The negative peak associated with the repolarization is less expressed.
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Parallel electrical measurements (Fig. 5, traces b–e and f–i) demonstrate a similar dependence of the AP time evolution on light/darkness. A transition of a cell from darkness to light prolonged the duration of the membrane depolarization for 2 s after a transitional period of 20–30 min on average. The depolarization time prolongation was accompanied by an increase in the depolarization peak of 10–15%. This can be attributed to the hyperpolarization of the resting potential under illumination (Mimura and Tazawa, 1986).
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t/to. Here, t is the light-induced delay with respect to the initial AC measured as the temporal shift in the depolarization maximum and to the time of the depolarization maximum for a particular illumination/darkness condition relative to the stimulus. We see that increases with the duration of illumination when cells are transferred from darkness into constant light. Fig. 7 summarizes the dependence of the average value of the normalized time delay as a function of illumination time obtained from AP measurements on 15 C. corallina internodal cells. Figs. 6 and 7 demonstrate the behavior of obtained from magnetic field (AC) and AP measurements of cells exposed to different durations of illumination. Both curves are almost identical. In addition, we also see that error bars in Fig. 7 are shorter (here were considered more cells).
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20 min, the magnetic signal intensity started to decrease in a way reminiscent of the behavior in the dark, as expected from our data in the illumination studies. Two cells were tested with both showing the same behavior.
Modeling
C. corallina internodal cells are, to a very good approximation, cylindrically symmetric and therefore ideally suitable for electrophysiological model studies. Several models followed the HH work on the nerve axon (Hodgkin and Huxley, 1952) and were adapted to C. corallina internodal cells (Beilby, 1982) or were devoted to studies of relations between AP, AC, and associated magnetic field (Clark and Plonsey, 1966; Woosley et al., 1985; Slibar et al., 2000). Here we will try to show that AP in C. corallina internodal cells experiences similarity to AP in those animal cells (heart, brain) where the "biochemical AP" is known to exist in addition to HH AP.
The HH type model is based purely on the kinetics of plasma membrane channels and the electrical properties of membrane but does not account for changes in [Ca2+]c. This factor, however, requires consideration in the context of this study because alterations in [Ca2+]c were observed experimentally under the influence of light/dark transitions (Miller and Sanders, 1987) and are in qualitative agreement with our model prediction. We have used an extended model of Othmer (Wacke et al., 2003), which describes the dynamics of [Ca2+]c in the context of a second messenger system. In that model, the AP in Chara is successfully described by an electrically stimulated release of Ca2+ from internal stores. The resulting changes in membrane conductance are the direct consequence of this dynamic change in [Ca2+]c. In this extended model, the thresholdlike dependence of Ca2+ mobilization on electrical stimulation can be simulated by combining the following two processes:
The model has been proved suitable to simulate a range of experimental results in the context of the AP in Chara, including a complex behavior of Ca2+ mobilization under periodic stimulation with higher-order phase locking and irregular responses upon increased stimulation frequency. This demonstrates that the all-or-none type activation of the action potential is only the consequence of the preceding all-or-none type mobilization of Ca2+ from internal stores. The dependency of the gating of the receptor channel on [Ca2+]c suggests that the Ca2+ concentration before stimulation of the AP has an influence on the following excitation kinetics.
To examine the effect of variable [Ca2+]c on the kinetics of the electrically stimulated changes in Ca2+, we modified the model as follows: Cells move excess Ca2+ from the cytoplasm back into internal stores by an endogenous Ca2+ pump system (e.g., Reddy, 2001). In the model, this process is accounted for by a Hill function:
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and 
are the Hill coefficients, and C is the cytosolic Ca2+ concentration. For more details, see Othmer (1997). To achieve different [Ca2+]c under resting conditions, which are known to occur during light/dark transitions (Miller and Sanders, 1987; Plieth et al., 1998a), we varied the Hill factor in Eq. 1. This procedure is only an indirect approach, since the chloroplasts from which the Ca2+ originates during light/dark transitions (Miller and Sanders, 1987) are not considered as extra Ca2+ pool in the model for [Ca2+]c dynamics. Nonetheless, this simple manipulation of the model should be sufficient to provide basic insight into the dependency of Ca2+ mobilization on [Ca2+]c.
Fig. 8 illustrates the results of this simulation. As a consequence of reduced Ca2+ pump activity, the resting [Ca2+]c increases over the physiological range from 20 nM to 200 nM. This roughly covers the changes in [Ca2+]c of 50–250 nM found in response to light/dark transitions in Chara (Miller and Sanders, 1987; Plieth et al., 1998a). The simulation further shows that an elevation of [Ca2+]c before the stimulation results in a progressive shortening of the delay time between stimulation and the rapid phase of [Ca2+]c rise. The dependence of this delay time on the resting [Ca2+]c concentration is plotted in the inset of Fig. 8.
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; Thiel et al., 1997), the model simulation compares well with the experimental data. The assumed progressive decrease in [Ca2+]c of 150 nM after the transition from dark to light results in an increasing delay time between the electrical stimulus and the rapid phase of [Ca2+]c rise, or the activation of the Cl– channels, respectively. |
ANALYSIS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSANALYSIS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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). This current reflects the summed contribution of all transmembrane ionic currents as well as electrogenic transport and other secondary transport processes in intracellular compartments. The SQUID microscope with its high spatial resolution will be a convenient instrument in studying these events in details. In those cases where different transmembrane ionic currents can be observed as transiently separable events, the analysis and the interpretation of the data can be undertaken. This is possible, for example, in the case of the influence of light on the kinetics of ionic currents during the AP. The main observation in this work was that a transfer of cells from the light into the dark resulted in an appreciable and reversible change in the kinetics of the action potential. Predominantly, the first part of the AC/AP temporal evolution, i.e., the slow depolarization part, increased progressively with time of light exposure; the rapid part of the AP, including depolarization and repolarization, was on the other hand only marginally affected. These observed changes in the kinetics of the Chara AP could in principle be modeled by a classical HH approach by adding a variable delay factor. However, there is no mechanistic motivation for such a delay factor, and it is difficult to envisage how a first-order voltage-dependent process could produce such a long and variable delay.
The experimental observations are, on the other hand, fully consistent with the view that the AP in Chara is of a "chemical" nature, i.e., based on a quasi all-or-none type liberation of Ca2+ from internal stores (Wacke and Thiel, 2001; Wacke et al., 2003). The model for the Chara AP (Wacke et al., 2003) predicts that a decrease of the resting [Ca2+]c by 250 nM, as is expected to occur in response to a dark/light transition (Miller and Sanders, 1987; Plieth, 1995), causes a delay in the onset of the rapid rise in [Ca2+]c during excitation. Assuming that the activation of the Cl– channels, which cause the depolarization, is the direct consequence of the change in [Ca2+]c, the measured data nicely match the model prediction, even on a quantitative basis. The model further predicts that a modulation of the resting [Ca2+]c has no big effect on the kinetics of the bulk changes of [Ca2+]c. Again, this prediction is met by the present data because the features in the temporal evolution of the AC/AP were basically unchanged.
Previous measurements have assessed the changes in [Ca2+]c after light/dark transitions indirectly by recording the velocity of cytoplasmic streaming, because this velocity is proportional to [Ca2+]c (Mccurdy and Harmon, 1992). In long-term recordings, Plieth and co-workers (Plieth et al., 1998b) found that it took 30 min to achieve a steady streaming velocity after transferring plants from dark to light. A slow time course for the change in Ca2+ in a similar order of magnitude can also be deduced from direct recordings of [Ca2+]c in Chara. The usual rise in [Ca2+]c after light/dark transitions was even after 5 min in the dark not yet fully saturated (Plieth, 1995). Hence, the time course in Ca2+ changes after light/dark transitions are of the same order of magnitude as the one we found in our measurements within experimental errors. Therefore, the entire observation of the change in the AP kinetics found under light/dark transition can be quantitatively explained by a slow progressing change in the resting [Ca2+]c and a consequent modulation of this parameter on the release of Ca2+ from internal stores.
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CONCLUSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSANALYSIS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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Our experimental results demonstrate the potential of SQUID microscope imaging for noninvasive magnetic measurements of axial intracellular current in electrically stimulated single internodal cells of Chara corallina under the influence of light. This kind of measurement provides a valuable technique to complement the longer existing electrophysiological methods in studies of plant cells.
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ACKNOWLEDGEMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSANALYSIS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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This research has been supported in part by a National Research Council Collaboration in Basic Science and Engineering program and by National Institutes of Health (NIH) grants R42 GM54963 and R01 HL58241. The SQUID microscope has been developed under NIH grant R42GM54963 and National Science Foundation grant DBI 9604948.
Submitted on April 23, 2004; accepted for publication October 13, 2004.
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