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First published online October 13, 2006; 10.1104/pp.106.087882 Plant Physiology 142:1460-1468 (2006) © 2006 American Society of Plant Biologists NAD(P)H Oscillates in Pollen Tubes and Is Correlated with Tip Growth1Department of Biology and Plant Biology Graduate Program, University of Massachusetts, Amherst, Massachusetts 01003 (J.G.K., P.K.H.); Departamento de Biologia Molecular de Plantas, Instituto de Biotecnologia, Universidad Nacional Autónoma de México, Morelos 62271, Mexico (L.C.); and Department of Biology, Long Island University, Brooklyn, New York 11201 (S.T.M.)
The location and changes in NAD(P)H have been monitored during oscillatory growth in pollen tubes of lily (Lilium formosanum) using the endogenous fluorescence of the reduced coenzyme (excitation, 360 nm; emission, >400 nm). The strongest signal resides 20 to 40 µm behind the apex where mitochondria (stained with Mitotracker Green) accumulate. Measurements at 3-s intervals reveal that NAD(P)H-dependent fluorescence oscillates during oscillatory growth. Cross-correlation analysis indicates that the peaks follow growth maxima by 7 to 11 s or 77° to 116°, whereas the troughs anticipate growth maxima by 5 to 10 s or 54° to 107°. We have focused on the troughs because they anticipate growth and are as strongly correlated with growth as the peaks. Analysis of the signal in 10-µm increments along the length of the tube indicates that the troughs are most advanced in the extreme apex. However, this signal moves basipetally as a wave, being in phase with growth rate oscillations at 50 to 60 µm from the apex. We suggest that the changes in fluorescence are due to an oscillation between the reduced (peaks) and oxidized (troughs) states of the coenzyme and that an increase in the oxidized state [NAD(P)+] may be coupled to the synthesis of ATP. We also show that diphenyleneiodonium, an inhibitor of NAD(P)H dehydrogenases, causes an increase in fluorescence and a decrease in tube growth. Finally, staining with 5-(and-6)-chloromethyl-2',7'-dichlorohydrofluorescein acetate indicates that reactive oxygen species are most abundant in the region where mitochondria accumulate and where NAD(P)H fluorescence is maximal.
Pollen tube growth, which is essential for sexual reproduction in higher plants, is highly polarized and extremely fast (approximately 200–300 nm/s in lily [Lilium formosanum]; Hepler et al., 2001  ). In our quest to decipher the mechanism of growth, we have taken advantage of the fact that the growth rate oscillates, with changes in lily occurring from 100 to 500 nm/s during a period of 20 to 50 s. Because many of the underlying physiological processes also oscillate with the same period but not usually with the same phase and amplitude as growth, we have examined phase relationships using cross-correlation analysis to determine if a process precedes or follows growth (Holdaway-Clarke and Hepler, 2003). Anticipatory events are of particular interest because they may bear a more direct, causal role than following events in the growth regulatory mechanism.
Among the oscillatory processes, most attention has been directed toward ions, with the conclusion that chloride efflux mirrors growth rate (Zonia et al., 2002
In this study, we focus on NAD(P)H in growing lily pollen tubes. These essential coenzymes occupy a central role in the control of cellular metabolism (Møller, 2001
NAD(P)H Colocalizes with Mitochondria When exciting at 360 nm and collecting the emission using a 400-nm long pass filter, we detect a fluorescent signal along the length of the pollen tube. Because these images are based on excitation and emission at single wavelengths, the amounts of fluorescence are not corrected for changes in optical path length. Nevertheless, by scanning a midline along the length of the tube, where the accessible volume will be relatively uniform, it becomes apparent that the signal is enriched in the apical 50 µm, with the strongest component occurring about 20 to 40 µm back from the apex, which corresponds to the base of the clear zone (Fig. 1, A and B ). To provide further information about associated structures, we stained living cells with Mitotracker Green, a fluorescent dye that labels mitochondria. It can be seen in Figure 1 (C and D) that mitochondria, which occur along the length of the pollen tube, tend to accumulate in the region 20 to 40 µm back from the apex. Line scan measurements along the length of the pollen tubes further indicate the close spatial localization of the NAD(P)H signal with the accumulation of mitochondria (Fig. 1, E and F).
NAD(P)H Fluorescence Oscillates during Pollen Tube Growth Oscillation Because pollen tubes of lily longer than 700 µm reliably exhibit oscillations in their growth rate, we selected cells at this state of growth for the analysis of NAD(P)H fluorescence. Pollen tubes were thus repetitively excited by 360 nm light for 250 ms at 3-s intervals over a total time period of 10 min. Images from a segment of sequential data are shown in Figure 2 ; a graphic display of representative oscillations is shown in Figure 3 . The resulting fluorescent signals clearly show distinct oscillations in which the peak fluorescence displays the same period as growth but not the same phase (Fig. 3). To provide more certainty on this point, we used autocorrelation to estimate the average periods of NAD(P)H titers and tip-growth rate within five independent pollen tubes; while the periods varied between 19 and 24.9 s between tubes, the periods of tip growth and NAD(P)H oscillation were virtually identical within tubes (data not shown). These oscillations in NAD(P)H fluorescence were not observed in tubes that did not exhibit growth oscillations, e.g. pollen tubes starting to grow or terminating growth. Here, we only observe a constant signal in the subapical region (data not shown). We also determined that these oscillations in signal were not due to a nonspecific effect, such as changes in accessible volume. Thus, single wavelength measurements from cells loaded with rhodamine dextran, which fills the accessible cytoplasm and does not respond to ions (calcium or pH), failed to show an oscillatory pattern (data not shown).
To determine the phase relationship with growth, we subjected these data to cross-correlation analysis (Fig. 4 ). The results show that in the tip of the pollen tube, the NAD(P)H fluorescent peaks follow growth maxima by 7 to 11 s or 77° to 116°. However, the troughs in the data are also thought to contain important information. It is known, for example, that the strongest fluorescent signal derives from protein-bound NAD(P)H, whereas free NAD(P)H has a substantially weaker fluorescent signal (Wakita et al., 1995 ; Paul and Schneckenburger, 1996; Blinova et al., 2005). The oscillation, therefore, could arise from a change between bound and free forms. Alternatively, there could be an oscillation between reduced and oxidized forms, where the latter [NAD(P)+] is nonfluorescent. Accordingly, we looked at the negative view of these fluorescent traces in cross-correlation analysis. The results indicate that in the apical-most domain, the troughs anticipate peak growth by 5 to 10 s or 54° to 107°. In addition, the cross-correlation analysis reveals that the troughs are as strongly correlated with growth as are the peaks.
To increase our understanding of the spatial nature of this signal, we examined fluorescence in 10-µm increments from a point centered at 5 µm from the tip to a point centered at 55 µm back using cross-correlation analysis of growth rate with regional NAD(P)H fluorescence (Fig. 4, B–G). In all areas, we are able to detect an oscillatory signal; however, the data show that the troughs are most advanced from growth at the extreme apex, with a gradual decline until 55 µm, when the troughs are in phase with the growth rate (Fig. 4H). A similar analysis of the peaks reveals that they progressively fall further behind growth. Because each subarea of NAD(P)H measurement is being cross correlated with the same set of tip growth data, it is also evident that the apical areas of NAD(P)H fluorescence are most highly correlated with tip growth (Fig. 4, B–G); however, the strength of cross correlation progressively attenuates and reaches approximately 60% of the peak correlation at 55 µm from the tip.
To modify the levels of NAD(P)H and see how this affects both fluorescence and tube growth, we applied diphenyleneiodonium (DPI), which inhibits plasma membrane NAD(P)H oxidases and mitochondrial NAD(P)H dehydrogenases and other flavoproteins (Roberts et al., 1995
Reactive Oxygen Species Production Correlates Spatially with Mitochondrial Distribution
In other systems, notably, root hairs (Foreman et al., 2003
This observation can be further confirmed from the line scan measurement, which indicates that the subapical region has an elevated level of ROS (Fig. 6C). Further studies involving a comparison of the ROS stain with Mitotracker Green reveal that the region with high ROS production correlates with the distribution of mitochondria and is largely excluded from the region of the inverted cone in the apex (compare Figs. 1D and 6B).
The results show that the amount of NAD(P)H, based on its endogenous fluorescence, spatially correlates with the distribution of mitochondria (Fig. 1). Not only do we recognize that mitochondria are the major source of NAD(P)H in the cell, but the studies showing that the components of the glycolytic pathway are also associated with outer membrane of the mitochondria (Giegé et al., 2003 ) ensure that additional enzymes associated with NAD(P)H metabolism will be similarly localized. Although oxidation of NAD(P)H occurs at the plasma membrane in soybean (Glycine max) hypocotyls and even exhibits an oscillatory profile (Morré and Morré, 1998; Morré, 2004), this process will be only a fraction (10%) of that generated by mitochondria. In addition, the oscillatory period for this process is 24 min and thus is 40 times longer than that reported herein in pollen tubes. We conclude, therefore, that mitochondrial metabolism accounts for the signal that we observe.
Of particular importance, the NAD(P)H-dependent fluorescence in pollen tubes oscillates and relates in a meaningful way to the corresponding oscillatory changes in the rate of growth (Figs. 2 and 3). When the signal is examined by cross-correlation analysis, the results indicate that the peak fluorescence follows the peak in growth rate. However, a similar analysis of the troughs indicates that they anticipate changes in growth rate (Fig. 4). A question that is often raised concerns how we can have confidence within an oscillatory context that an event either anticipates or follows changes in growth rate. To resolve this issue, we have used cross-correlation analysis, which permits us to establish the strength of covariance and lag between two processes, e.g. tip growth and process "x" (Brillinger, 1975
Although the cross-correlation analysis indicates that the troughs and peaks are equal in their strength of correlation to growth, we focus on the troughs because they indicate a process that anticipates the changes in growth rate. It is our view that a preceding or anticipatory event may bear a closer relationship to the central regulator of growth than one that follows. The troughs represent a decrease in NAD(P)H fluorescence, which could be due to two different processes. First, it could be due to a shift from bound to free NAD(P)H in which the bound form exhibits a much stronger fluorescence than that which is free (Wakita et al., 1995
Of additional interest, the troughs are most advanced over growth in the apical domain, but they become progressively less advanced in more basipetal regions until they are in phase with growth rate at about 50 to 60 µm back from the tip (Fig. 4). Moreover, the troughs show a stronger correlation to growth rate in the apical domains where they are most advanced over growth rate. If the troughs represent a relative increase in NAD(P)+, then the data indicate that the production of this oxidized species occurs first in the extreme apex and sets up a wave that spreads basipetally. The rearward propagation of the NAD(P)+ wave may represent the recharging of the coenzyme using resources that are posterior to the tip. Mitochondria, while lower in number relative to the subapical region, are nevertheless present very close to the tip (Fig. 1D; Lancelle and Hepler, 1992
Mitochondria extending close to the apex will also experience an oscillating calcium gradient (Holdaway-Clarke and Hepler, 2003
It is attractive to envision that the oxidation of NAD(P)H, yielding NAD(P)+, is directly coupled to the synthesis of ATP, which is then used to power a variety of processes localized to the pollen tube apex. There could by many key steps that require ATP; however, we draw attention to a few events that are confined to the apical 10 µm of the pollen tube. First, we note the internal alkaline band (Feijó et al., 1999
Additional results from this study underscore the relationship between NAD(P)H and growth, and emphasize the importance of achieving the correct balance between the oxidized and reduced forms of these coenzymes. Thus, the inhibition of NAD(P)H dehydrogenase activity with DPI leads to an increase in NAD(P)H and eventually to a decrease in the growth rate (Fig. 5). We also provide preliminary evidence about ROS production in pollen tubes. In agreement with several others (Pei et al., 2000 In conclusion, we have shown that NAD(P)H fluorescence oscillates during oscillatory pollen tube growth. Mathematical analysis of the phase relationship between NAD(P)H fluorescence and growth rate indicates that at least one component, possibly NAD(P)+, anticipates the changes in growth rate and might therefore occupy a position close to the central growth regulator. We favor the idea that oxidation of NAD(P)H with the concomitant production of ATP constitutes a primal event in pollen tube growth control. We also recognize that the NAD(P)H oxidation may be associated with formation of ROS and these may serve an important signaling function.
Pollen Tube Growth Pollen grains of lily (Lilium formosanum) were germinated and grown in lily growth medium (LGM), which is composed of 15 mM MES, 1.6 mM H3BO3, 1 mM KCl, 0.1 mM CaCl2, and 7% (0.2 M) Suc at pH 5.5. Pollen tubes were plated on a coverslip in an open glass slide chamber by mixing 35 µL of LGM containing the recently germinated pollen tubes and 35 µL of LGM, which contained 1.4% of low gelling point agarose (type VII; Sigma). This step turned out to be very critical as the layer of agarose on the coverslip should be as thin as possible to keep the pollen tubes flat and as close to the coverslip as possible. To ensure a thin layer, the coverslips were previously treated with hydrofluoric acid (5%) for 10 min, which slightly etched the glass and facilitated the tight binding of the agarose layer.
NAD(P)H is fluorescent and has been visualized by exciting at 340 to 365 nm and detecting emission at 400 to 500 nm (Lakowicz et al., 1992 Following experimentation with different filters on our microscope system, we found the best signal with the following setup: 360 nm (10-nm bandpass) as excitation filter, 380 nm dichroic, and 400-nm long pass emission filter (all filters from Chroma). Under these circumstances, the signal is evident and could be followed in living pollen tubes. Nevertheless, the signal is weak; therefore, we employed binning (2 x 2), which markedly increased the signal-to-noise ratio and allowed us to decrease the exposure time to 250 ms. These conditions permitted us to obtain a measurement every 3 s and thus to record fluorescence changes at multiple points during 5 to 10 min of oscillatory growth. Because the above measurements of NAD(P)H were made at a single wavelength, we felt it was important to rule out nonspecific factors that could affect the signal. In particular, we were concerned about oscillatory changes in accessible volume because we had found in related studies that the endoplasmic reticulum displays a periodic movement in the apical domain during oscillatory pollen tube growth. To test for oscillations in accessible volume, we loaded pollen tubes with rhodamine dextran, an inert, space-filling dye, and measured its fluorescence changes in different regions of the tube during oscillatory growth.
For mitochondrial staining, we used Mitotracker Green (Molecular Probes), which was dissolved in LGM at a final concentration of 2.5 µM. The pollen tubes were stained in the dye medium for 15 min, after which this medium was removed. The cells were rinsed once in LGM without dye and cultured in fresh LGM. Excitation was carried out at 495 nm, with fluorescence emission at 516 nm. To detect ROS, we used CM-H2DCFDA (Molecular Probes). The stock dye was prepared at 20 µM in dimethyl sulfoxide. A total of 50 µL of this stock solution was applied to the growth chamber containing the pollen tubes for 20 min. Afterward, the cells were rinsed with dye-free medium and observed in the microscope. To obtain information about the relative changes in amount of ROS, we microinjected a reference dye (70 kD tetramethylrhodamine dextran; Molecular Probes) into pollen tubes previously loaded with CM-H2DCFDA. Image pairs were then acquired; first, we excited CM-H2DCFDA at 480 nm and collected the emission at 530 nm (ROS dependent), and, second, we excited tetramethyl-rhodamine dextran (70 kD) at 555 nm and collected the emission at 600 nm (ROS independent). A ratio of these two signals permitted us to map out the relative levels of ROS in the pollen tube.
DPI is a general flavin dehydrogenase inhibitor; it has been used as an inhibitor of NAD(P)H oxidation (Roberts et al., 1995
Many physical and chemical phenomena in the pollen tube tip are convoluted in the four dimensions of space and time. We take a series of measurements as two-dimensional differential interference contrast (DIC) and fluorescence images and can isolate phenomena in those images as regions of pixels that identify the growing tip or an absorbance or fluorescence measure of a chemical. These localized phenomena may be oscillating with varying periods and amplitudes, i.e. they are convoluted in ways the naked eye may not be able to discern. Deconvolution of the phenomena involves taking the recorded localized oscillations and mathematically reconvolving them in a pair-wise fashion to decide whether particular phenomena are changing with their own program, or if they are in concert with or phase-offset from another tube tip phenomenon. In this way, we are able to order the timing and spatial relations of events that occur in the tube tip. Data were collected for growth and NAD(P)H fluorescence in 100 to 200 3-s bins with a DIC image plus a fluorescence image in each bin. The DIC image, taken in 50 ms, produced the tip location, and those from adjacent bins were used to calculate growth rate, which was assigned to the mid-time between two successive DIC images. The fluorescence image was taken over a 1-s interval and assigned to a time midway through its collection. Fluorescence integrations were made in nonoverlapping, 10-µm increments along the length of the tube.
For long-term effects on growth rate and fluorescence to be minimized, the data were normalized and smoothed (Cleveland, 1981
We used cross-correlation analysis (Brillinger, 1975
The convolve function (Brillinger, 1975
We thank Professor Max Møller, from The Royal Veterinary and Agricultural University, Frederiksberg, Denmark, for many helpful comments during the preparation of this study. We also thank our colleagues at University of Massachusetts for stimulating discussions. Received August 16, 2006; accepted October 10, 2006; published October 13, 2006.
1 This work was supported by the National Science Foundation (grant nos. MCB–0055799 and MCB–0516852 to P.K.H.) and by Dirección General de Asuntos para el Personal Académico/Universidad Nacional Autónoma de México (grant no. IN–228903 and postdoctoral research to L.C.).  The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Luis Cárdenas (luisc{at}ibt.unam.mx). www.plantphysiol.org/cgi/doi/10.1104/pp.106.087882 * Corresponding author; e-mail luisc{at}ibt.unam.mx; fax 52–7773136600.
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