We thank Prof. Geoffrey Wasteneys, University of British Columbia, for seed-expressing RFP-TUB6 and an anonymous reviewer for helpful corrections. We are grateful to Prof. Hugh Dickinson for alerting us to the use of cellulose film as a mechanical support and to John Baker for photography. This work was supported by BBSRC research grants BB/G013993/1 and BB/D004055/1 to I.M., and a BBSRC Doctoral Training Award and Clarendon Scholarship to C.K.

1. Creating the Chamber
<ol>
	<li>Using a glass cutter, cut 3 mm wide strips from the end of a 1 mm thick microscope slide. Using a cyanoacrylate super-glue or double-sided tape, glue these glass strips approximately 45 mm apart across the width of a second microscope slide (Figure 1A). One slide per chamber is required, plus additional slides to pour agar slabs (one slide will yield 2-3 agar slabs). Slides can be reused.</li>
	<li>Between the glass strips, fashion a gasket of identical height out of gas-permeant poly(dimethylsiloxane) gum. Place a ball of poly(dimethylsiloxane) gum onto the slide (Figure 1B), wet a second slide with a small quantity of 100% absolute ethanol, and flatten the poly(dimethylsiloxane) gum ball with the second slide until it has reached the height of the glass strips (Figure 1C).
	<ol>
		<li>If necessary, trim excess poly(dimethylsiloxane) gum with a razor blade and repeat flattening.</li>
	</ol>
	</li>
	<li>Using a suitable cutter or razor blade wetted in absolute ethanol, remove the interior part of the poly(dimethylsiloxane) gum to create the cavity that will later hold the seedling and agar slab (Figure 1D).</li>
	<li>Carefully trim the gel with a razor blade to create a gasket with a final wall thickness of approximately 2 mm (Figure 1E). The permeability of gases through a poly(dimethylsilaxane) barrier is independent of thicknesses above 50 &#181;m11.</li>
</ol>
2. Creating the Agar-solidified Slab of Growth Medium
<ol>
	<li>On a microscopy slide with two glass strips, place a coverslip (22 mm x 50 mm) so that it rests on both glass strips.</li>
	<li>Pipette melted half-strength Murashige and Skoog growth medium containing 1% w/v sucrose and 1.5% w/v agar (&#189; MS) into the resulting space below the coverslip until the latter is completely filled (approximately 1 mL total volume) and leave until agar is set (approximately 5 min). 
	Note: Pharmacological compounds can be administered through the agar slab by adding appropriate concentrations to the liquid medium before the slab is poured. This was successfully tested for dexamethasone12, isoxaben, 2,6-dichlorobenzonitrile (DCB), oryzalin, and latrunculin B.</li>
</ol>
3. Finishing the Chamber Setup
<ol>
	<li>Air equilibrate PFD by shaking a small volume of PFD in a tube7. Add a small amount (approximately 200 &#181;L) of air-equilibrated PFD to the well of the gel gasket, but do not fill the chamber completely. This PFD will help to avoid the trapping of air bubbles under the agar slab as it is placed into the chamber.</li>
	<li>Remove the coverslip from the agar slab (at step 2.2 above) and use a razor blade to cut off a portion of desired size and shape. Then place it into the well of the gel gasket (Figure 1F). There should be a gap of 2-4 mm to the gasket all around.</li>
	<li>Fill the chamber completely with air-equilibrated PFD.</li>
	<li>Place one or more A. thaliana seedlings (up to 3 seedlings for imaging over 2-3 days) onto the agar slab with the cotyledons and hypocotyl hanging over the edge, floating in PFD (Figure 1G).</li>
	<li>To obtain seedlings, sterilize seeds with 70% ethanol, plant on &#189; MS medium supplemented with 1% sucrose and 0.8% agar, stratify for 2-4 days at 4 &#176;C, and grow on vertically oriented plates at 22 &#176;C in a 16 h light/8 h dark regime. For imaging lateral roots, grow plants for 7-10 days before transferring to imaging chambers.</li>
	<li>To close the chamber, apply a coverslip matched to the optics of the objective, gently pressing it down with the edge of a glass microscope slide until the coverslip rests on both glass strips (Figure 1H).</li>
	<li>Secure the coverslip with strips of 1.25 cm wide micropore surgical tape cut in half length-wise on each end if desired (Figure 1I). Allow the specimen to settle for approximately 30 min before imaging. This minimizes specimen movement during imaging.</li>
	<li>Perform imaging with an upright microscope to maintain a controlled substrate for growth. Use 20X/0.7NA or 63X/1.2NA CS2 objectives.</li>
</ol>

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R., Love, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simple+method+for+imaging+Arabidopsis+leaves+using+perfluorodecalin+as+an+infiltrative+imaging+medium.">A simple method for imaging Arabidopsis leaves using perfluorodecalin as an infiltrative imaging medium.</a> J Vis Exp. , (2012).</li><li>Littlejohn, G. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+update:+improvements+in+imaging+perfluorocarbon-mounted+plant+leaves+with+implications+for+studies+of+plant+pathology,+physiology,+development+and+cell+biology.">An update: improvements in imaging perfluorocarbon-mounted plant leaves with implications for studies of plant pathology, physiology, development and cell biology.</a> Front Plant Sci. 5, 140 (2014).</li><li>Dias, A., Freire, M., Coutinho, J. 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The authors have nothing to disclose.

The method presented here is a simple strategy for high-resolution confocal imaging of developing lateral roots for two and up to three days. For periods of up to 48 h, no adverse effects of the imaging system on lateral root development were observed. After 48 h, the average lateral root growth began to slow, although a substantial subset of the roots (37%) continued to grow at rates comparable to average root growth on plates. Therefore, through imaging a sufficiently large number of roots, roots whose growth slows after 48 h can be excluded. Systematic tests of the imaging chambers were not performed for periods longer than 72 h, but alternative strategies are recommended if extended imaging periods are desired. Imaging chambers may be left on the microscope stage continuously, if suitable environmental conditions are provided, or removed to a growth facility and periodically returned to the microscope. This allows numerous chambers to be imaged in parallel.
One advantage of the chambers described here is that lateral roots are fixed in their position and can be imaged using high-resolution water immersion lenses. The spatial stability critically depends on the agar concentration used in the supporting agar slab. Initially a range of different concentrations from 0.8% agar to 2% agar were tested, revealing that high concentrations in this range stably fixed roots in space, but root growth slowed down faster and some roots exhibited signs of mechanical stress, including reduced cell elongation. By contrast, low agar concentrations did not provide the required support and roots drifted in x, y, and z during the imaging. The optimal 1.5% agar slab fixes the position of the specimen without adverse mechanical effects. Under these conditions, after settling in the first 30 min or so, roots are stable over many hours, permitting overnight data acquisition. During acquisition of 4D data, z-stack ranges were typically bracketed by an additional 5-10 &#181;m but this was principally to accommodate out-of-plane growth of lateral roots rather than z-drift or wobble. Although the standard agar concentration provides some resistance against penetration, gravitropically growing roots will eventually penetrate the agar. However, through a minor modification of the imaging chamber, root growth could be maintained parallel to the coverslip, allowing older lateral roots and primary roots to be imaged. Furthermore, the basic imaging chamber could easily be customized for hypocotyls. Hypocotyls are more freely floating in this system so the z-axis bracket for image acquisition was increased usually to about 20 &#181;m. In this study, an upright microscope was used throughout, which allowed the substrate properties to be controlled. The imaging chamber may be adaptable to inverted microscope configurations but the time-dependent influence of the rigid coverslip on intercepting organs will need to be evaluated.
Littlejohn and colleagues have pointed out that PFD itself does not readily dissolve biological molecules, which means it cannot be used for the directly for the delivery of pharmacological compounds7. This problem was overcome by supplying such compounds through the slab of solidified growth medium on which the agar slab rests. While perfusion systems will remain the method of choice for wash-out experiments, the imaging chamber has been used successfully for the administration of dexamethasone12 and other compounds. One note, while this article was in preparation von Wagenheim and colleagues18 described a chamber for imaging lateral root development using light-sheet microscopy.

To study cellular and subcellular processes that underlie plant development, there is an increasing demand for high-resolution long-term time-lapse imaging strategies. A key challenge in such imaging experiments is the maintenance of physiological conditions including sufficient gaseous exchange plus a supply of water and nutrients1,2,3. To utilize objectives with high numerical apertures for optimal optical resolution, specimens should be positioned in close proximity and oriented parallel to the coverslip. Movement in x, y, and z directions should ideally be minimal during imaging.
While seedlings are often mounted in water or aqueous solution for short-term imaging, water has a low capacity for dissolving CO2 and O2 (1.54 mg/mL and 0.04 mg/mL, respectively at 20 &#176;C, 0.1 MPa)4, which makes it unsuitable for extended time-lapse experiments. Perfusion systems that maintain constant levels of oxygen and nutrients are one solution to this problem and have been developed for both confocal laser scanning microscopy (CLSM)1,2,3 and light sheet microscopy (LSM)5. Systems like the RootChip2 and the RootArray3 have been designed specifically for time lapse imaging of developing roots and involve germinating seeds in a custom-build multi-specimen device. These arrangements ensure minimal mechanical perturbation and are designed for the parallel analysis of multiple seedlings, but are not optimized for imaging of subcellular structures. Calder and colleagues have designed a more complicated perfusion-based imaging chamber optimized for imaging of subcellular structures in which the specimen is held in position by a mesh to allow the use of high magnification immersion lenses1.
Presented here is an alternative, simple solution to this problem which is not based on perfusion systems, but utilizes perfluorodecalin (PFD), a perfluorocarbon that has recently gained popularity as a mounting medium for Arabidopsis imaging6,7,8. In such applications, the high capacity of PFD for dissolving CO2 and O2 (1.9 g/mL for O2 in PFD compared to 0.04 mg/mL in water)9, allows gaseous exchange by the immersed tissue. Furthermore, PFD is non-fluorescent and its refractive index (1.313) is comparable with that of water (1.333) and is closer to that of cytosol (~1.4) than air (1.000)6. Perfluorocarbons have been reported to have minimal physiological effect on a variety of plants and plant tissues6, with radish seeds germinating readily when submersed in PFD and exhibiting normal growth and development for at least two full days when supplied with water10. Similar observations have been made for germinating Arabidopsis seeds6. Based on stimulated Raman scattering to directly image the distribution of PFD in Arabidopsis leaf tissues after infiltration, Littlejohn and colleagues conclude that PFD is likely not taken up by living cells8. PFD has previously been used predominantly to image aerial tissues, where it significantly increases imaging depth as it readily infiltrates air spaces due to its low surface tension6. Here, PFD is adopted for long-term confocal imaging of developing lateral roots. In this configuration, one or more seedlings are placed onto a slab of growth medium solidified with agar and immersed in PFD. The PFD allows gaseous exchange in the imaging chamber, preventing anoxia. PFD is highly volatile so it is retained by a gasket of poly(dimethylsiloxane) gum that also has high gas permeability (1.5 x 10-12 pmol m-1 s-1 Pa-1 for CO2)11. Nutrients and water are supplied by the slab of medium solidified with agar. At the same time, this agar slab gently presses the root against the coverslip, thus fixing its relative position in the imaging chamber and allowing the use of high-resolution water immersion lenses. Furthermore, the agar slab can be used to administer a range of pharmacological treatments, including dexamethasone, oryzalin, and isoxaben. The imaging chambers can be assembled in large numbers from standard microscopy slides using minimal equipment. The imaging chambers were developed and characterized to study lateral root development but are adaptable to imaging other seedling organs such as primary root tips and hypocotyls.

<table><tbody><tr><td>Perfluorodecalin</td><td>F2 Chemicals, F2 Chemicals Ltd., Lea Town, Lancashire, UK</td><td>FLUTEC PP6</td><td></td></tr><tr><td>Poly(dimethylsiloxane) gum&amp;nbsp;</td><td>Carolina Biological Supply; Burlington, NC, USA</td><td>Item # 132700</td><td>Carolina Observation Gel</td></tr><tr><td>Cavity Microscope Slides</td><td>VWR International Ltd, Lutterworth,&amp;nbsp;UK</td><td>10118-600</td><td>Cavity is 13 mm diameter and 0.2-0.4 mm in depth. Any cavity slide will probably suffice</td></tr><tr><td>Cyanoacrylate glue</td><td>Loctite</td><td>604753</td><td>Any &#039;super-glue&#039; suitable for glass will probably suffice</td></tr><tr><td>Cellulosic cellophane membrane</td><td>AA Packaging Limited, Preston, UK</td><td></td><td>325P cellulose film; 80 mm disc</td></tr></tbody></table>

a simple chamber for long-term confocal imaging of root and hypocotyl development

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and subcellular processes, high resolution time-lapse microscopy strategies that preserve physiological conditions are required. Plant tissues must have adequate nutrient and water supply with sustained gaseous exchange but, when submerged and immobilized under a coverslip, they are particularly susceptible to anoxia. One strategy that has been successfully employed is the use of a perfusion system to maintain a constant supply of oxygen and nutrients. However, such arrangements can be complicated, cumbersome, and require specialized equipment. Presented here is an alternative strategy for a simple imaging system using perfluorodecalin as an immersion medium. This system is easy to set up, requires minimal equipment, and is easily mounted on a microscope stage, allowing several imaging chambers to be set up and imaged in parallel. In this system, lateral root growth rates are indistinguishable from growth rates under standard conditions on agar plates for the first two days, and lateral root growth continues at reduced rates for at least another day. Plant tissues are supplied with nutrients via an agar slab that can be used also to administer a range of pharmacological compounds. The system was established to monitor lateral root development but is readily adaptable to image other plant organs such as hypocotyls and primary roots.

Lateral roots grow at physiological rates in imaging chambers. 
Lateral root lengths of plants in imaging chambers were measured at hourly intervals (Figure 2A, left, Movie 1, n = 23) and growth rates were compared to those of lateral roots grown on standard Petri plates containing the same agar-solidified medium (Figure 2A, right, Movie 2, n = 23). Growth rates can vary considerably between lateral roots, with the length of the root being one determining factor owing to the increasingly longer zones of growth and proliferation13. Therefore, sets of lateral roots were selected to represent roots of different initial lengths (ranging from 50 &#181;m to 1150 &#181;m) in both the imaging chamber and the agar plate. Average lateral root length at the start of the experiment was similar in each set (439 and 442 &#181;m for imaging chambers and plates, respectively). In the analyzed time interval of 27 h, lateral root growth was approximately linear both in imaging chambers and on plates, with an average growth rate of 52 &#181;m/h (R2 = 0.997) on plates and 51 &#181;m/h in imaging chambers (R2 = 0.993) (Figure 2B). Growth rates of individual lateral roots where highly variable both on plates and in imaging chambers (Figure 2C), ranging from 17 &#181;m/h to 82 &#181;m/h in imaging chambers and from 13 &#181;m/h to 87 &#181;m/h on plates. While growth rates generally increased with the length of the root, growth rates still varied substantially between roots of similar length but the variance was similar in imaging chambers and on plates.
Lateral roots proliferate in imaging chambers and can be imaged using high numerical-aperture objectives. 
Lateral roots co-expressing the plasma-membrane marker NPSN12-YFP14 and the microtubule marker UBQ1::mRFP-TUBULIN BETA6 (RFP-TUB6)15 were imaged by CLSM10 at 1 h intervals to document microtubule behavior during cell proliferation in imaging chambers (Figure 2D, Movie 3). In these chambers, lateral roots were highly stable in their position on the x, y and z axes, allowing the acquisition of continuous time-lapse confocal stacks over several hours. Meristematic cells continuously proliferated, as is evident from the formation of preprophase bands (Figure 2D, green arrows), mitotic spindles (Figure 2D, white arrows), and phragmoplasts (Figure 2D, blue arrows). As with seedlings on Petri plates, fully elongated root cells in imaging chambers initiated root hairs (Figure 2A, Figure 2D, arrow heads), indicating further that development proceeded as expected in the imaging chambers. To provide a wide field-of-view, the images in Figure 2D were obtained with a 20x/0.7 NA objective. It was also possible to obtain high resolution images of lateral roots using a high numerical-aperture 63x/1.2 NA water-immersion objective (Figure 2E).
Lateral root growth is sustained in imaging chambers for up to three days.
To explore how long imaging chambers could support lateral root growth at physiological rates, lateral root length in imaging chambers (n = 27) and on plates (n = 33) was quantified at 0 h, 24 h, 48 h, and 72 h. The roots chosen were shorter than 200 &#181;m at 0 h, with an average lateral root length of 117 &#181;m in imaging chambers and 121 &#181;m on plates. Since shorter roots grow more slowly on average (Figure 2C), they grew over the edge of the agar slab less often, and were easier to image. The average growth of all lateral roots was not significantly different in imaging chambers compared to Petri plates in the first 48 h (Figure 3A). However, the average growth was significantly reduced between 48 h and 72 h (Figure 3A). While growth rates appear to generally slow down in imaging chambers after 48 h, growth rates between individuals remain highly variable (Figure 3B, 3C), which means that there is a substantial subset of lateral roots that grows at comparable rates on plates and in chambers over the complete 72 h growth period. 23 of the 33 lateral roots grown on plates (69.7%) reach a length within one standard deviation of the mean length at 72 h (between 927 and 3,163 &#181;m, Figure 3B, red). 10 of 27 (37.0%) lateral roots in imaging chambers reach a length within this interval (Figure 3B, red).
Imaging chambers can easily be adapted for older lateral roots, primary roots and hypocotyls.
One of the limitations of the original imaging chamber design was that while the stiff agar slab provided some mechanical resistance, gravitropically growing roots eventually penetrated the agar slab, resulting in loss of image quality (Figure 4B) and growth beyond the working distance of high NA lenses, even the relatively long working-distance (0.3 mm) 63X/1.3 NA CS2 objective. Young lateral roots lack the statolith-containing columella cells required for gravitropism17 so they initially grew parallel to the coverslip in imaging chambers (Figure 2E). As they grow longer and columella and statoliths form, lateral roots exhibit increasing positive gravitropism, while primary roots exhibit a strong gravitropic response from germination onwards. This restricts to a few hours the period over which primary roots can be imaged and it severely limits the starting-length of lateral roots that can be imaged continuously for 48 h or more. To overcome this limitation, the chamber was modified such that growth is maintained parallel to the coverslip by a cellulosic cellophane membrane that acts as a penetration barrier on the surface of the agar slab (Figure 4A). Initial attempts using a 1.5% agar slab caused reduced root growth within the first 24 h, owing perhaps to mechanical stress. Lower agar concentrations in the slab were tested and it was found that in chambers with a slab containing 0.8% agar and cellulose film, young lateral root growth rates did not differ significantly from growth rates in conventional chambers for the first 48 h. For 0-24 h, mean root growth in cellulose chambers and conventional chambers was 242 &#181;m and 262 &#181;m respectively (p = 0.78 Student&#39;s t-Test) and for 24-48 h was 330 &#181;m and 355 &#181;m1 respectively (p = 0.67 Student&#39;s t-Test); n = 12 and n = 11, respectively. This arrangement effectively prevented lateral roots from growing away from the coverslip, into the agar, and also allowed imaging of primary roots for up to 48 h (Figure 4C).
To test whether the imaging chambers could be adapted for organs other than roots, Arabidopsis hypocotyls were chosen. Hypocotyls are substantially thicker than roots so in the conventional imaging chamber, the uppermost cells were squeezed against the coverslip, leading to deformation. An alternative design was therefore developed using cavity slides that contain an oval depression in their center (Figure 4D). In this configuration, a 1 mm thick slab of agar (prepared as in 2.2 above) was positioned with one end protruding by a few mm into the slide cavity (Figure 4D). Hypocotyls were positioned above the cavity in the slide while the root was positioned above the horizontal part. This ensured that the seedling was fixed in space by the root, but the hypocotyl was not squashed. While growth rates in chambers compared to plates were not systematically quantified, hypocotyls in imaging chambers exhibited the well-described wave of longitudinal growth migrating up the hypocotyl (Figure 4E, 4F)16.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/55331/55331fig1.jpg" /> 
Figure 1: Assembling an imaging chamber. All details are described in the text. <a href="http://cloudflare.jove.com/files/ftp_upload/55331/55331fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/55331/55331fig2.jpg" /> 
Figure 2: Lateral roots grow under physiological conditions in imaging chambers. (A) Arabidopsis lateral root development in an imaging chamber (left) and on a plate (right) over 25 h (constant light, horizontal incubation, simultaneous imaging). Scale bars = 200 &#181;m. (B) Plot showing lateral root lengths over 27 h (measured in 1 h intervals), of lateral roots as shown in (A). Individual roots plotted in grey (n = 23 for plates and imaging chambers), average length as red circles. Error bars = 1 SD. Red line is linear fit through average lengths. (C) Plot showing average growth rate for all lateral roots shown in (B) relative to their initial length. (D) Maximum intensity projections of 3D confocal stacks acquired from lateral roots expressing NPSN12-YFP and RFP-TUB6 at consecutive time points. Inset: Magnification of boxed region. Green arrows: preprophase bands; white arrows: mitotic spindles; blue arrows: phragmoplasts; arrowheads: root hairs. (E) Maximum intensity projections of high-resolution images acquired from a young lateral root of the transgenic line shown in (D) using a 63X/1.2 NA objective and Nyquist sampling (pixel dimensions 77 nm x 77 nm) at 0 h and 16 h; asterisks locate identical cells at each time point; arrow indicates a cell that divides between 0h and 16 h. Scale bars = 50 &#181;m (A-C); 20 &#181;m (E). <a href="http://cloudflare.jove.com/files/ftp_upload/55331/55331fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/55331/55331fig3.jpg" /> 
Figure 3: Long-term lateral root development in imaging chambers. (A) Plot showing average absolute root growth in three consecutive 24 h intervals in imaging chambers compared to plates. n.s. indicates p &#62;0.05; *** indicates p &#60;0.001 (student t-test). n = 33 (plates) and n = 27 (imaging chambers). (B) Plot showing lateral root lengths at consecutive time points on all roots used in (A). Red: all lateral roots with a final length within 1 SD of the mean of all lateral roots grown on plates (between 927 and 3163 &#181;m). (C) Maximum intensity projections of representative 3D confocal stacks of the plasma-membrane marker NPSN12-YFP in lateral roots, acquired at consecutive time points over 72 h. Scale bars = 100 &#181;m. <a href="http://cloudflare.jove.com/files/ftp_upload/55331/55331fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/55331/55331fig4.jpg" /> 
Figure 4: Imaging chambers can be adapted for other plant organs. (A-C) Imaging chamber adaption for primary roots. (A) Schematic of imaging chamber design. This is identical to the standard design (Figure 1) apart from two modifications: the MS medium slab contains 0.8% agar instead of 1.5% agar, and a cellulose film (red) is placed between agar and plant to prevent growth into the agar. (B) XY (top) and XZ (bottom) maximum intensity projections of representative 3D confocal stacks of the plasma-membrane marker NPSN12-YFP in the primary root of a 7 day old seedling imaged at 0 h and 48 h in a conventional imaging chamber. Note that the root grows into the agar due to its gravitropic response. (C) XY (top) and XZ (bottom) maximum intensity projections of representative 3D confocal stacks of the plasma-membrane marker NPSN12-YFP in the primary root of a seven-day old seedling imaged at 0 h and 48 h in the imaging chamber with cellulose film. Before application, the cellulose film was sterilized in 80% ethanol and soaked in liquid &#189; MS medium. Note that gravitropic growth of the root into the agar is prevented. (D-F) Imaging chamber adaptation for hypocotyls. (D) Schematic showing hypocotyl imaging chamber design. A poly(dimethylsiloxane) gum gasket (grey) is manufactured on a cavity slide between two glass strips (yellow). A slab of 1.5% agar of even thickness (beige) is placed on the slide, partially reaching into the cavity. The chamber is filled with PFD (blue). Seedlings are placed onto the agar slab so that the hypocotyl occupies the downwards-sloping region of the agar slab in the cavity while the root is positioned the horizontal part of the agar. The chamber is closed with a coverslip. (E) Maximum intensity projections of representative 3D confocal stacks of the plasma-membrane marker NPSN12-YFP in a hypocotyl of a two-day old seedling imaged over 48 h at consecutive time points. (F) Longitudinal growth in two consecutive time intervals of individual cells along the basal to apical axis (relative positions at 0 h along the axis shown in E) of the hypocotyl shown in (E). Note the previously described wave of growth migrating up the hypocotyl16. Scale bars = 100 &#181;m. <a href="http://cloudflare.jove.com/files/ftp_upload/55331/55331fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
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Watch this Scientific Journal Video about A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development at JoVE.com

A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

Presented here is a simple technique for high-resolution confocal time-lapse imaging of root and hypocotyl development for up to 3 days using high numerical-aperture objectives and perfluorodecalin as an immersion medium.

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and ...

a-simple-chamber-for-long-term-confocal-imaging-root-hypocotyl

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

10.1K Views.  University of Oxford.  Hier wordt een simpele techniek voor de confocal time-lapse beeldvorming van wortel- en hypocotyloplossing met hoge resolutie voor maximaal 3 dagen gegeven met behulp van hoge numerieke diafragma-doelstellingen en perfluorodecaline als onderdompelingsmedium. 

Video: A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration have been thoroughly analyzed in vitro. However, in vivo cell migration somehow differs from in vitro migration, and has proven more difficult to analyze, being less accessible to direct observation and manipulation. This protocol uses the migration of the prospective prechordal plate in the early zebrafish embryo as a model system to study the function of candidate genes in cell migration. Prechordal plate progenitors form a group of cells which, during gastrulation, undergoes a directed migration from the embryonic organizer to the animal pole of the embryo. The proposed protocol uses cell transplantation to create mosaic embryos. This offers the combined advantages of labeling isolated cells, which is key to good imaging, and of limiting gain/loss of function effects to the observed cells, hence ensuring cell-autonomous effects. We describe here how we assessed the function of the TORC2 component Sin1 in cell migration, but the protocol can be used to analyze the function of any candidate gene in controlling cell migration in vivo.

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Combining cell transplantation, cytoskeletal labeling and loss/gain of function approaches, this protocol describes how the migrating zebrafish prospective prechordal plate can be used to analyze the function of a candidate gene in in vivo cell migration.

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration ...

Time-lapse Fluorescence Imaging of Arabidopsis Root Growth with Rapid Manipulation of The Root Environment Using The RootChip

The root functions as the physical anchor of the plant and is the organ responsible for uptake of water and mineral nutrients such as nitrogen, phosphorus, sulfate and trace elements that plants acquire from the soil. If we want to develop sustainable approaches to producing high crop yield, we need to better understand how the root develops, takes up a wide spectrum of nutrients, and interacts with symbiotic and pathogenic organisms. To accomplish these goals, we need to be able to explore roots in microscopic detail over time periods ranging from minutes to days.
We developed the RootChip, a polydimethylsiloxane (PDMS)- based microfluidic device, which allows us to grow and image roots from Arabidopsis seedlings while avoiding any physical stress to roots during preparation for imaging1 (Figure 1). The device contains a bifurcated channel structure featuring micromechanical valves to guide the fluid flow from solution inlets to each of the eight observation chambers2. This perfusion system allows the root microenvironment to be controlled and modified with precision and speed. The volume of the chambers is approximately 400 nl, thus requiring only minimal amounts of test solution.
Here we provide a detailed protocol for studying root biology on the RootChip using imaging-based approaches with real time resolution. Roots can be analyzed over several days using time lapse microscopy. Roots can be perfused with nutrient solutions or inhibitors, and up to eight seedlings can be analyzed in parallel. This system has the potential for a wide range of applications, including analysis of root growth in the presence or absence of chemicals, fluorescence-based analysis of gene expression, and the analysis of biosensors, e.g. FRET nanosensors3.

This article provides a protocol for cultivation of Arabidopsis seedlings in the RootChip, a microfluidic imaging platform that combines automated control of growth conditions with microscopic root monitoring and FRET-based measurement of intracellular metabolite levels.

The root functions as the physical anchor of the plant and is the organ responsible for uptake of water and mineral nutrients such as nitrogen, ...

Bioengineering

Live Confocal Imaging of Developing Arabidopsis Flowers

The study of plant growth and development has long relied on experimental techniques using dead, fixed tissues and lacking proper cellular resolution. Recent advances in confocal microscopy, combined with the development of numerous fluorophores, have overcome these issues and opened the possibility to study the expression of several genes simultaneously, with a good cellular resolution, in live samples. Live confocal imaging provides plant biologists with a powerful tool to study development, and has been extensively used to study root growth and the formation of lateral organs on the flanks of the shoot apical meristem. However, it has not been widely applied to the study of flower development, in part due to challenges that are specific to imaging flowers, such as the sepals that grow over the flower meristem, and filter out the fluorescence from underlying tissues. Here, we present a detailed protocol to perform live confocal imaging on live, developing Arabidopsis flower buds, using either an upright or an inverted microscope.

Live confocal imaging provides biologists with a powerful tool to study development. Here, we present a detailed protocol for the live confocal imaging of developing Arabidopsis flowers.

The study of plant growth and development has long relied on experimental techniques using dead, fixed tissues and lacking proper cellular resolution. ...

Confocal Live Imaging of Shoot Apical Meristems from Different Plant Species

The shoot apical meristem (SAM) functions as a conserved stem cell reservoir and it generates almost all aboveground tissues during the postembryonic development. The activity and morphology of SAMs determine important agronomic traits, such as shoot architecture, size and number of reproductive organs, and most importantly, grain yield. Here, we provide a detailed protocol for analyzing both the surface morphology and the internal cellular structure of the living SAMs from different species through laser scanning confocal microscope. The whole procedure from the sample preparation to the acquisition of high resolution three-dimensional (3D) images can be accomplished within as short as 20 minutes. We demonstrate that this protocol is highly efficient for studying not only the inflorescence SAMs of the model species but also the vegetative meristems from different crops, providing a simple but powerful tool to study the organization and development of meristems across different plant species.

This protocol presents how to live image and analyze the shoot apical meristems from different plant species using laser scanning confocal microscopy.

The shoot apical meristem (SAM) functions as a conserved stem cell reservoir and it generates almost all aboveground tissues during the postembryonic ...

Long-term Time Lapse Imaging of Mouse Cochlear Explants

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building the highly ordered, complex structure of the mammalian cochlea proceeds for around ten days. In order to study changes in gene expression over this period and their response to pharmaceutical or genetic manipulation, long-term imaging is necessary. Previously, live imaging has typically been limited by the viability of explanted tissue in a humidified chamber atop a standard microscope. Difficulty in maintaining optimal conditions for culture growth with regard to humidity and temperature has placed limits on the length of imaging experiments. A microscope integrated into a modified tissue culture incubator provides an excellent environment for long term-live imaging. In this method we demonstrate how to establish embryonic mouse cochlear explants and how to use an incubator microscope to conduct time lapse imaging using both bright field and fluorescent microscopy to examine the behavior of a typical embryonic day (E) 13 cochlear explant and Sox2, a marker of the prosensory cells of the cochlea, over 5 days.

Live imaging of the embryonic mammalian cochlea is challenging because the developmental processes at hand operate on a temporal gradient over ten days. Here we present a method for culturing and then imaging embryonic cochlear explant tissue taken from a fluorescent reporter mouse over five days.

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building ...

Neuroscience

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by their natural environment, root hair morphology and function are difficult to study and often excluded from plant research. In recent years, microfluidic platforms have offered a way to visualize root systems at high resolution without disturbing the roots during transfer to an imaging system. The microfluidic platform presented here builds on previous plant-on-a-chip research by incorporating a two-layer device to confine the Arabidopsis thaliana main root to the same optical plane as the root hairs. This design enables the quantification of root hairs on a cellular and organelle level and also prevents z-axis drifting during the addition of experimental treatments. We describe how to store the devices in a contained and hydrated environment, without the need for fluidic pumps, while maintaining a gnotobiotic environment for the seedling. After the optical imaging experiment, the device may be disassembled and used as a substrate for atomic force or scanning electron microscopy while keeping fine root structures intact.

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

This article demonstrates how to culture Arabidopsis thaliana seedlings in a two-layer microfluidic platform that confines the main root and root hairs to a single optical plane. This platform can be used for real-time optical imaging of fine root morphology as well as for high-resolution imaging by other means.

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by ...

Development of Microfluidic Devices to Study the Elongation Capability of Tip-growing Plant Cells in Extremely Small Spaces

In vivo, tip-growing plant cells need to overcome a series of physical barriers; however, researchers lack the methodology to visualize cellular behavior in such restrictive conditions. To address this issue, we have developed growth chambers for tip-growing plant cells that contain a series of narrow, micro-fabricated gaps (~1 &#181;m) in a poly-dimethylsiloxane (PDMS) substrate. This transparent material allows the user to monitor tip elongation processes in individual cells during microgap penetration by time-lapse imaging. Using this experimental platform, we observed morphological changes in pollen tubes as they penetrated the microgap. We captured the dynamic changes in the shape of a fluorescently labeled vegetative nucleus and sperm cells in a pollen tube during this process. Furthermore, we demonstrated the capability of root hairs and moss protonemata to penetrate the 1 &#181;m gap. This in vitro platform can be used to study how individual cells respond to physically constrained spaces and may provide insights into tip-growth mechanisms.

We describe a method to investigate the capability of tip-growing plant cells, including pollen tubes, root hairs, and moss protonemata, to elongate through extremely narrow gaps (~1 &#181;m) in a microfluidic device.

In vivo, tip-growing plant cells need to overcome a series of physical barriers; however, researchers lack the methodology to visualize cellular behavior ...

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue patterning. During animal development, key cell proliferation and patterning events occur very quickly. For instance, in Caenorhabditis elegans all cell divisions required for the larval body plan are completed within six hours after fertilization, with seven mitotic cycles1; the sixteen or more mitoses of Drosophila embryogenesis occur in less than 24 hr2. In contrast, cell divisions during plant development are slow, typically on the order of a day 3,4,5 . This imposes a unique challenge and a need for long-term live imaging for documenting dynamic behaviors of cell division and differentiation events during plant organogenesis. Arabidopsis epidermis is an excellent model system for investigating signaling, cell fate, and development in plants. In the cotyledon, this tissue consists of air- and water-resistant pavement cells interspersed with evenly distributed stomata, valves that open and close to control gas exchange and water loss. Proper spacing of these stomata is critical to their function, and their development follows a sequence of asymmetric division and cell differentiation steps to produce the organized epidermis (Fig. 1).
This protocol allows observation of cells and proteins in the epidermis over several days of development. This time frame enables precise documentation of stem-cell divisions and differentiation of epidermal cells, including stomata and epidermal pavement cells. Fluorescent proteins can be fused to proteins of interest to assess their dynamics during cell division and differentiation processes. This technique allows us to understand the localization of a novel protein, POLAR6, during the proliferation stage of stomatal-lineage cells in the Arabidopsis cotyledon epidermis, where it is expressed in cells preceding asymmetric division events and moves to a characteristic area of the cell cortex shortly before division occurs. Images can be registered and streamlined video easily produced using public domain software to visualize dynamic protein localization and cell types as they change over time.

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

We describe a protocol using chamber slides and media to immobilize plant cotyledons for confocal imaging of the epidermis over several days of development, documenting stomatal differentiation. Fluorophore-tagged proteins can be tracked dynamically by expression and subcellular localization, increasing understanding of their possible roles during cell division and cell-type differentiation.

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue ...

Biology

Measuring Spatial and Temporal Ca2+ Signals in Arabidopsis Plants

Developmental and environmental cues induce Ca2+ fluctuations in plant cells. Stimulus-specific spatial-temporal Ca2+ patterns are sensed by cellular Ca2+ binding proteins that initiate Ca2+ signaling cascades. However, we still know little about how stimulus specific Ca2+ signals are generated. The specificity of a Ca2+ signal may be attributed to the sophisticated regulation of the activities of Ca2+ channels and/or transporters in response to a given stimulus. To identify these cellular components and understand their functions, it is crucial to use systems that allow a sensitive and robust recording of Ca2+ signals at both the tissue and cellular levels. Genetically encoded Ca2+ indicators that are targeted to different cellular compartments have provided a platform for live cell confocal imaging of cellular Ca2+ signals. Here we describe instructions for the use of two Ca2+ detection systems: aequorin based FAS (film adhesive seedlings) luminescence Ca2+ imaging and case12 based live cell confocal fluorescence Ca2+ imaging. Luminescence imaging using the FAS system provides a simple, robust and sensitive detection of spatial and temporal Ca2+ signals at the tissue level, while live cell confocal imaging using Case12 provides simultaneous detection of cytosolic and nuclear Ca2+ signals at a high resolution.

Measuring Spatial and Temporal Ca2+ Signals in Arabidopsis Plants

Ca2+ signaling regulates diverse biological processes in plants. Here we present approaches for monitoring abiotic stress induced spatial and temporal Ca2+ signals in Arabidopsis cells and tissues using the genetically encoded Ca2+ indicators Aequorin or Case12.

Developmental and environmental cues induce Ca2+ fluctuations in plant cells. Stimulus-specific spatial-temporal Ca2+ patterns are sensed by cellular Ca2+ ...

Light Sheet Fluorescence Microscopy of Plant Roots Growing on the Surface of a Gel

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the observation of the whole organ with a high spatial- as well as temporal resolution over prolonged periods of time, which may cause photo-toxic effects. This protocol shows a plant sample preparation method for light-sheet microscopy, which is characterized by mounting the plant vertically on the surface of a gel. The plant is mounted in such a way that the roots are submerged in a liquid medium while the leaves remain in the air. In order to ensure photosynthetic activity of the plant, a custom-made lighting system illuminates the leaves. To keep the roots in darkness the water surface is covered with sheets of black plastic foil. This method allows long-term imaging of plant organ development in standardized conditions.

This protocol shows a plant sample preparation method for light-sheet microscopy. The setup is characterized by mounting the plant vertically on the surface of a gel and letting it grow in controlled bright conditions. This allows long-term observation of plant organ development in standardized conditions.

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the ...

A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

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