Name: measurement of leaf hydraulic conductance and stomatal conductance and their responses to irradiance and dehydration using the evaporative flux method (efm)
Uploaded: 2012-12-31T12:00:00
Duration: 12 min 11 s
Description: Watch this Scientific Journal Video about Measurement of Leaf Hydraulic Conductance and Stomatal Conductance and Their Responses to Irradiance and Dehydration Using the Evaporative Flux Method EFM at

We are grateful to four anonymous reviewers for improving the manuscript, and to the National Science Foundation (Grant IOS-0546784) for support.

1. Measuring Leaf Hydraulic Conductance for Hydrated Leaves (Kmax)
<ol>
 <li>Collect shoots preferably in the evening or night such that xylem tensions are low, cut in air and place in dark plastic bag filled with wet paper towels. In the lab, recut at least two nodes from the end of every shoot under water. Let shoot rehydrate overnight in pure water, or other solution to be used for Kleaf measurements. Ultrapure water is recommended as a flow solution unless the impacts of different solutions are to be investigated as a factor.</li>
 <li>Cut the leaf from the shoot with a fresh razor blade under partially degassed flow solution. Ultrapure water is recommended as a flow solution unless the impacts of different solutions are to be investigated as a factor. We recommend degassing the solution overnight in a flask using a vacuum pump, but other methods may be employed.</li>
 <li>Wrap pre-stretched parafilm around the petiole to ensure a good seal between petiole and tubing. This step is especially helpful for non-round petioles, as wrapping extra parafilm will help round the petiole to fit inside the tubing. Dental paste can be applied to the petiole before wrapping the parafilm to fill grooves in petioles.</li>
 <li>Connect the leaf to silicone tubing under ultrapure water to prevent air entering the system. As the leaf is lifted above the source of water, the water in the tubing is under subatmospheric pressure, and thus the seal is good if no air is drawn into the tubing during the measurement. Alternatively to sealing the petiole directly into silicone tubing, a compression fitting with rubber gasket can be used (e.g., (Omnifit A2227 bore adaptor; Omnifit, Cambridge, UK); this approach is useful for petioles with especially complex shapes, or for grass blades, which can be wrapped around a glass rod and sealed with the compression fitting. When feasible, sealing the petiole directly into silicone tubing has the advantage of being rapid and leaving a petiole-tubing junction that is transparent, allowing bubbles to be more easily seen. The tubing connects the leaf to hard tubing running to a graduated cylinder on a balance (&plusmn;10 &mu;g resolution), which logs data every 30 sec to a computer (e.g. using "Balance Link" Software; Mettler-Toledo GmbH, Greifensee, Switzerland) for the calculation of flow rate through the leaf.</li>
 <li>Leaves should be held adaxial surface upwards in wood frames strung with fishing line above a large box fan and under a light source ( &gt; 1,000 &mu;mol &bull; m2 &bull; s-1 photosynthetically active radiation for high-irradiance measurements), slightly above the level of the meniscus in the graduated cylinder. The leaf should be placed above the water in the graduated cylinder to ensure that water moving to the leaf is driven by transpiration, and not entering the leaf under pressure due to gravity, as would be the case if the leaf were placed below the meniscus of the water in the cylinder. Small containers filled with wet paper towel are placed inside the balance chamber to ensure the air in the balance is water-saturated, thus avoiding evaporation from the water source to the atmosphere in the balance. This possibility can be checked by halting flow into the leaf using a four-way stopcock valve in the tubing, and checking that the mass of the water cylinder on the balance does not decline.</li>
 <li>Place a clear Pyrex container filled with water above the leaf to absorb the heat of the lamp.</li>
 <li>Maintain leaf temperature between 23 &deg;C and 28 &deg;C throughout the experiment by adding more cold water to the Pyrex container such that the heat lamp does not heat the Pyrex container sufficiently to warm the air around the leaf below.</li>
 <li>Allow the leaf to transpire on the apparatus for at least 30 min, thus giving sufficient time to acclimate to high irradiance, and additionally until flow rate has stabilized, with no upward or downward trend, and with a coefficient of variation &lt;5% for at least 5-10 measurements made at 30 sec flow intervals. It is essential for the flow rate to reach a steady state because this method assumes a stable &Psi;leaf. Previous studies found these criteria to be sufficient for stabilization of flow rate (E), &Psi;leaf and Kleaf; tests with longer measurement periods after stable flow was established showed no relationship of Kleaf to measurement time for each of seven species of a wide range of leaf capacitance17,18 . Stabilization generally takes 30 min or less, but in some species may require more than an hour for some species.</li>
</ol>
Note: When flow rate is very low (&lt;8 &mu;g s-1), stability can be determined using the running average of the last five 30 sec intervals.
<ol start="9">
 <li>Discard measurements if the flow suddenly changes, either due to sudden stomatal closure, or apparent leakage from the seal or blockage in the system by particles or air bubbles.</li>
 <li>When flow stabilizes, record leaf temperature with a thermocouple.</li>
 <li>Quickly remove the leaf from the tubing, dab dry the petiole and place the leaf into a sealable bag which had been previously exhaled in, to halt transpiration.</li>
 <li>Let the leaf equilibrate in its bag for at least 20 min.</li>
 <li>Average the final 10 flow rate measurements. This will be your E measurement.</li>
 <li>When the leaf is equilibrated, measure the final leaf water potential (&Psi;final) with a pressure chamber.</li>
 <li>Measure leaf area with a flatbed scanner and image processing software, or with a leaf area meter.</li>
 <li>Kleaf is calculated as E / -&Delta;&Psi;leaf (where &Delta;&Psi;leaf = &Psi;final - 0 MPa) and further normalized by leaf area. Units are in mmol m-2 s-1 MPa-1. Because the leaf is positioned just above the level of water in the graduated cylinder, the effect of gravity in reducing the pressure of water entering the leaf would be negligible, especially relative to the &Psi;final, which is always &gt; -0.1 MPa.</li>
 <li>To correct for changes in Kleaf induced by temperature dependence of water viscosity15,19,20 , standardize Kleaf values to 25 &deg;C by using the following equation:</li>
</ol>
<img alt="Equation 1" fo:content-width="3.863in" fo:src="/files/ftp_upload/4179/4179eq1highres.jpg" src="/files/ftp_upload/4179/4179eq1.jpg" />
<ol start="18">
 <li>
Stomatal conductance (gs) can be determined as E divided by the leaf-to-air vapor pressure difference, equivalent to the vapor pressure deficit (VPD), determined using a temperature and relative humidity sensor adjacent to the leaf. VPD is calculated using the following equation21</li>
</ol>
<img alt="Equation 2" src="/files/ftp_upload/4179/4179eq2.jpg" fo:content-width="1.67in" fo:src="/files/ftp_upload/4179/4179eq2highres.jpg" /> 
where RH is the relative humidity, VP is the saturation vapor pressure of the air (in kPa; a function of temperature, according to the Arden Buck equation22), and AP the atmospheric pressure (in kPa).
2. Measuring Leaf Hydraulic Conductance for Dehydrated Leaves
<ol>
 <li>Collect shoots preferably in the evening or night such that xylem tensions are low, cut in air and place in dark plastic bags filled with wet paper towels. In the lab, recut at least two nodes from the end of every shoot under water. Let shoot rehydrate overnight in pure water, or other solution to be used for Kleaf measurements. Ultrapure water is recommended as a flow solution unless the impacts of different solutions are to be investigated as a factor.</li>
 <li>Cut shoots into segments with at least three leaves under ultrapure water and dehydrate shoots with a fan for different periods of time to a range of &Psi;leaf values.</li>
 <li>Once shoots are dehydrated, place sealable bags (Whirl-Pak; Nasco, Fort Atkinson, WI, USA) which have been previously exhaled in) around each leaf of the shoot, and then place the whole shoot in a large sealable bag with wet paper towel.</li>
 <li>Let the shoot equilibrate for at least 30 min (for strongly dehydrated shoots longer equilibration times may be necessary).</li>
 <li>After equilibration, excise the top and bottom leaf and measure initial &Psi;leaf (=&Psi;o) using a pressure chamber.</li>
 <li>Discard the shoot if the difference in the &Psi;leaf of those two leaves is greater than 0.1 MPa (for strongly dehydrated leaves, this range can be extended to 0.3 MPa).</li>
 <li>The third leaf is then used to determine Kleaf and gs with the EFM, following steps 1.2 -1.18 above. Notably, steady state E can be obtained even for leaves that have been severely dehydrated leaves, as stomata still open and generate gs values substantially higher than cuticular conductance17.</li>
 <li>To construct hydraulic vulnerability curves, calculate Kleaf using the &Psi;final as in step 1.16, and then plot Kleaf values against whichever is lowest, &Psi;o or &Psi;final, i.e., the strongest dehydration experienced by the leaf during the experiment (=&Psi;lowest). Similarly, the decline of gs in response to leaf dehydration can be determined by plotting gs against &Psi;lowest. Previous work has shown that dehydration induces at least partially irreversible declines in Kleaf and gs14,16.</li>
 <li>We recommend obtaining at least 6 Kleaf values per 0.5 MPa interval of &Psi;lowest (this interval can be reduced to 0.25 MPa if the species you are working with is really vulnerable to dehydration).</li>
 <li>Perform Dixon outlier test on Kleaf values for each 0.5 MPa water potential interval of &Psi;lowest to remove outliers from your vulnerability curve23.</li>
 <li>Because species differ in their responses of Kleaf and gs to dehydration14,16, fit different functions for each species and select the maximum likelihood (lowest AIC value; see 14 for details). We recommend fitting at least four functions previously used in the literature: linear (<img alt="Equation 3" src="/files/ftp_upload/4179/4179eq3.jpg" fo:content-width="1.4in" fo:src="/files/ftp_upload/4179/4179eq3highres.jpg"/>), sigmoidal (<img alt="Equation 4" src="/files/ftp_upload/4179/4179eq4.jpg" fo:content-width="1.483in" fo:src="/files/ftp_upload/4179/4179eq4highres.jpg"/>), logistic (<img alt="Equation 5" src="/files/ftp_upload/4179/4179eq5.jpg" fo:content-width="1.7in" fo:src="/files/ftp_upload/4179/4179eq5highres.jpg"/>) and exponential (<img alt="Equation 6" src="/files/ftp_upload/4179/4179eq6.jpg" fo:content-width="1.583in" fo:src="/files/ftp_upload/4179/4179eq6highres.jpg"/>).</li>
</ol>
3. Representative Results
The decline of leaf hydraulic conductance with dehydration varies strongly across species (Fig. 1). Drought sensitive species experience a stronger decline at less negative water potentials than more drought tolerant species, when measured under high irradiance (PAR &gt; 1,000 &mu;mol &bull; m-2 &bull; s-1). The water potential at 80% loss of hydraulic conductance (P80) for mesic herb Helianthus annuus was 3 MPa less negative than that of Heteromeles arbutifolia, native to California chaparral. Heteromeles arbutifolia responded linearly to the decline in water potential, whereas Helianthus annuus showed a non-linear decline.
Both stomatal conductance (gs) and leaf hydraulic conductance (Kleaf) can also respond dramatically to irradiance (Fig. 2). For Raphiolepis indica, the stomatal conductance declined strongly with dehydration for leaves measured under high irradiance (PAR &gt; 1,000 &mu;mol &bull; m-2 &bull; s-1) and under lab light (PAR &lt; 6 &mu;mol &bull; m-2 &bull; s-1). Under high irradiance, leaves measured with the EFM that had been dehydrated to the water potential that is typical for well-lit leaves on intact plants at mid-day showed similar gs as for leaves measured on intact plants with a porometer (star in Fig. 2). Notably, for leaves at maximum hydration, Kleaf was four-fold higher for leaves exposed to high than low irradiance (11 vs 2.8 mmol m-2 s-1 MPa-1), and therefore Kleaf showed a stronger light response in well hydrated leaves than gs. Further, under high irradiance, Kleaf declined more strongly with dehydration. The P80 was 0.93 MPa less negative under high than low irradiance.
<img alt="Figure 1" src="/files/ftp_upload/4179/4179fig1.jpg" /> 
 Figure 1. Leaf hydraulic vulnerability curves for a drought sensitive (top) and a drought tolerant (bottom) species (from 14,24 ). Each point represents a single measurement of leaf hydraulic conductance (Kleaf) on a detached leaf using the evaporative flux method (EFM). The vertical bars represent the water potential at which the species lost 80% of initial leaf hydraulic conductance (P80). Best fit functions using maximum likelihood are plotted for both species.
<img alt="Equation 7" src="/files/ftp_upload/4179/4179eq7.jpg" fo:content-width="3.293in" fo:src="/files/ftp_upload/4179/4179eq7highres.jpg"/>
<img alt="Figure 2" src="/files/ftp_upload/4179/4179fig2.jpg" /> 
 Figure 2. The response of stomatal conductance (gs) and of leaf hydraulic conductance (Kleaf) to declining leaf water potential for Raphiolepis indica measured under high vs low irradiance (from 16). (PAR &gt; 1,000 and &lt; 6 mmol &bull; m-2 &bull; s-1). Each point represents a single measurement on a detached leaf using the evaporative flux method (EFM), with white and black dots representing leaves measured under high and low irradiance respectively. Equations for stomatal conductance curves in high vs low irradiance: <img alt="Equation 8" src="/files/ftp_upload/4179/4179eq8.jpg" fo:content-width="4.727in" fo:src="/files/ftp_upload/4179/4179eq8highres.jpg" /> and <img alt="Equation 9" src="/files/ftp_upload/4179/4179eq9.jpg" fo:content-width="2.26in" fo:src="/files/ftp_upload/4179/4179eq9highres.jpg"/>. Equations for vulnerability curves in high vs. low irradiance: <img alt="Equation 10" src="/files/ftp_upload/4179/4179eq10.jpg" fo:content-width="2.7in" fo:src="/files/ftp_upload/4179/4179eq10highres.jpg"/> and <img alt="Equation 11" src="/files/ftp_upload/4179/4179eq11.jpg" fo:content-width="1.8in" fo:src="/files/ftp_upload/4179/4179eq11highres.jpg"/>. The grey star in the upper panel represents the mean &plusmn; standard error for gs and leaf water potential measured on leaves of intact plants at mid-day with a porometer. Grey and black vertical bars in the lower panel represent the water potential at which the species lost 80% of initial leaf hydraulic conductance (P80) under high and low irradiance respectively.

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W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaporation+sites+and+distillation+in+leaves.">Evaporation sites and distillation in leaves.</a> Ann. Bot. 41, 1081-1082 (1977).</li><li>Canny, M. J., Huang, C. X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Leaf+water+content+and+palisade+cell+size.">Leaf water content and palisade cell size.</a> New Phytol. 170, 75-85 (2006).</li><li>Santiago, L. S., Mulkey, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+test+of+gas+exchange+measurements+on+excised+canopy+branches+of+ten+tropical+tree+species.">A test of gas exchange measurements on excised canopy branches of ten tropical tree species.</a> Photosynthetica. 41, 343-347 (2003).</li><li>Sack, L., Cowan, P. D., Jaikumar, N., Holbrook, N. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+'hydrology'+of+leaves:+co-ordination+of+structure+and+function+in+temperate+woody+species.">The 'hydrology' of leaves: co-ordination of structure and function in temperate woody species.</a> Plant Cell Environ. 26, 1343-1356 (2003).</li></ol>

No conflicts of interest declared.

The evaporative flux method presented here allows relatively rapid (30 min) determination of leaf hydraulic conductance in the laboratory with simultaneous measurement of stomatal conductance.
The EFM is so far the method that most closely follows the natural pathway of water in leaves, given that water evaporates in the leaf airspaces and diffuses from the stomata15. A number of other experimental methods for Kleaf determination exist25, with three being especially common. (1) In the high pressure flow method (HPFM), water is pushed through the leaf under high pressure15,26 . However, vulnerability curves cannot be obtained with this method because applied positive pressures may refill emboli and rehydrate mesophyll tissue. (2) In the rehydration kinetics method (RKM), Kleaf is calculated using the analogy between the rehydration of dehydrated leaves with the charging of a capacitor in series with a resistors 27,28. This method is rapid, but does not reproduce the entire system of natural transpirational pathways in the leaf and is not easily modified to allow measurement of leaves acclimated to high irradiance. (3) In the vacuum pump method (VPM) the leaf is placed in a chamber, its petiole connected to a water source on a balance, while vacuums of different intensities are applied to the leaf driving water loss from the leaf15,26 . This method can be time consuming. Measurements using all three methods have been found to give consistent results15,28,29 , and thus the EFM is frequently used given its being based on transpirational water movement, and its relative rapidity, and its modification for measuring Kleaf under different conditions.
All of the typical methods used for determining Kleaf involve a level of uncertainty in quantifying accurately the driving force corresponding to transpiration25. In the EFM, the &Delta;&Psi;leaf used to calculate Kleaf is lower than the true driving force, as it is based on the bulk &Psi;leaf after the leaf is equilibrated for the pressure bomb. At this point, the bulk &Psi;leaf does not necessarily represent the water potential of the cells at the end of the hydraulic pathway, where the water evaporates, but rather a volume-weighted average for the relatively few cells of low water potential where water is moving and evaporating, and for the more numerous cells of high water potential that may be isolated from the transpiration stream. How much the &Psi;leaf differs from the true driving force thus depends on where the water is evaporating in the leaf. If water evaporates throughout the leaf, the bulk &Psi;leaf may be close to the true driving force, and if water evaporates only from relatively few cells near the stomata, the the bulk &Psi;leaf may substantially underestimate the true driving force, i.e., the water potential of those cells may in fact be much lower, and thus actual Kleaf would be lower than that determined with the EFM. Thus, the degree that Kleaf determined by the EFM represents the true Kleaf is sensitive to the water flow pathways, which are still not well understood. While the EFM provides data that match those of other methods for measuring Kleaf (see above), and is excellent for comparative use, the possibility of species-variation in water flow pathways through the mesophyll needs further investigation. Indeed, even for leaves of given species, these water pathways might vary under different conditions, e.g., under different irradiances and heat loads4,30 or of different water status (given tissue shrinkage)31. Further knowledge of the water transport pathways would thus allow the EFM to be used to resolve the effects of the changing pathways on overall leaf hydraulic transport.
The measurement of stomatal conductance in the EFM has the advantages of allowing determination of responses to irradiance and dehydration, and providing matched responses of gs with Kleaf. Additionally, the effect of previous leaf dehydration followed by rehydration on gs can be assessed with this method. However, this method has two potential drawbacks. First, this gs measurement is made on an excised leaf, rather than on an intact plant, and second, the determination of stomatal conductance is based on climate measurement some distance from the leaf, rather than close to the leaf as in a porometer chamber, though the ventilation of leaf with the fan equilibrates the leaf surface with surrounding air. For some species, excised leaves may show different stomatal responses than for leaves on an intact plant 16,32 . However, for several species gs estimated using the EFM was similar to values measured on intact plants with a porometer for leaves at mid-day &Psi;leaf16 (Fig. 2).
The EFM can be applied for physiological insight, i.e., investigating the dynamics of Kleaf, its basis in structure and anatomy, and its responses to environmental factors. Additionally, the method can be used to compare plants of given species grown under different conditions, or of different ages, or to compare plants of different species adapted to different environments.
By constructing leaf vulnerability curves, one can gain insight into the mechanisms of species tolerances to drought. During drought, cavitation and cell collapse have been shown to occur in the leaf xylem of some species, and cell shrinkage in the mesophyll during dehydration could also impact water movement (reviewed in 14). Future studies will be needed to determine the relative importance of each of these processes in driving the decline of Kleaf and stomatal closure.
Irradiance is also a strong factor influencing Kleaf and the EFM is highly suited for measurements under high irradiance. The combined response of aquaporins to irradiance and drought should gain much interest. High irradiance leads to increased expression and/or activation of aquaporins that permit faster water flow through living cells, increasing Kleaf, whereas dehydration leads to reduced expression and/or de-activatation of aquaporins, reducing Kleaf. The combined impacts of irradiance and water supply on Kleaf require further investigation4,10,16,33 .
Further, across species, Kleaf and its vulnerability has been related to species distributions with respect to the supply of light water and other resources3,13,24 . Species from dry areas tend to show greater resistance to hydraulic decline, facing more negative soil water potentials, and by maintaining a high Kleaf at more negative water potentials, they can maintain high gs and continue capturing CO2 for photosynthesis13,14. Thus, the study of leaf hydraulic properties provides insights at the level of cell, leaf, and whole plant and its responses to the environment, and is likely to yield numerous important new discoveries at all scales.

measurement of leaf hydraulic conductance and stomatal conductance and their responses to irradiance and dehydration using the evaporative flux method (efm)

Water is a key resource, and the plant water transport system sets limits on maximum growth and drought tolerance. When plants open their stomata to achieve a high stomatal conductance (gs) to capture CO2 for photosynthesis, water is lost by transpiration1,2. Water evaporating from the airspaces is replaced from cell walls, in turn drawing water from the xylem of leaf veins, in turn drawing from xylem in the stems and roots. As water is pulled through the system, it experiences hydraulic resistance, creating tension throughout the system and a low leaf water potential (&Psi;leaf). The leaf itself is a critical bottleneck in the whole plant system, accounting for on average 30% of the plant hydraulic resistance3. Leaf hydraulic conductance (Kleaf = 1/ leaf hydraulic resistance) is the ratio of the water flow rate to the water potential gradient across the leaf, and summarizes the behavior of a complex system: water moves through the petiole and through several orders of veins, exits into the bundle sheath and passes through or around mesophyll cells before evaporating into the airspace and being transpired from the stomata. Kleaf is of strong interest as an important physiological trait to compare species, quantifying the effectiveness of the leaf structure and physiology for water transport, and a key variable to investigate for its relationship to variation in structure (e.g., in leaf venation architecture) and its impacts on photosynthetic gas exchange. Further, Kleaf responds strongly to the internal and external leaf environment3. Kleaf can increase dramatically with irradiance apparently due to changes in the expression and activation of aquaporins, the proteins involved in water transport through membranes4, and Kleaf declines strongly during drought, due to cavitation and/or collapse of xylem conduits, and/or loss of permeability in the extra-xylem tissues due to mesophyll and bundle sheath cell shrinkage or aquaporin deactivation5-10. Because Kleaf can constrain gs and photosynthetic rate across species in well watered conditions and during drought, and thus limit whole-plant performance they may possibly determine species distributions especially as droughts increase in frequency and severity11-14.
We present a simple method for simultaneous determination of Kleaf and gs on excised leaves. A transpiring leaf is connected by its petiole to tubing running to a water source on a balance. The loss of water from the balance is recorded to calculate the flow rate through the leaf. When steady state transpiration (E, mmol &bull; m-2 &bull; s-1) is reached, gs is determined by dividing by vapor pressure deficit, and Kleaf by dividing by the water potential driving force determined using a pressure chamber (Kleaf= E /- &Delta;&psi;leaf, MPa)15.
This method can be used to assess Kleaf responses to different irradiances and the vulnerability of Kleaf to dehydration14,16,17.

Watch this Scientific Journal Video about Measurement of Leaf Hydraulic Conductance and Stomatal Conductance and Their Responses to Irradiance and Dehydration Using the Evaporative Flux Method EFM at 

Measurement of Leaf Hydraulic Conductance and Stomatal Conductance and Their Responses to Irradiance and Dehydration Using the Evaporative Flux Method EFM

We describe a relatively rapid (30 min) and realistic method for simultaneously measurement of leaf hydraulic conductance (Kleaf) and stomatal conductance (gs) for transpiring excised leaves. The method can be modified to measure the light and dehydration responses of Kleaf and gs.

Measurement of Leaf Hydraulic Conductance and Stomatal Conductance and Their Responses to Irradiance and Dehydration Using the Evaporative Flux Method (EFM)

Water is a key resource, and the plant water transport system sets limits on maximum growth and drought tolerance. When plants open their stomata to ...

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Research

JoVE Journal

Biology

Stable Isotopic Profiling of Intermediary Metabolic Flux in Developing and Adult Stage Caenorhabditis elegans

Stable isotopic profiling has long permitted sensitive investigations of the metabolic consequences of genetic mutations and/or pharmacologic therapies in cellular and mammalian models. Here, we describe detailed methods to perform stable isotopic profiling of intermediary metabolism and metabolic flux in the nematode, Caenorhabditis elegans. Methods are described for profiling whole worm free amino acids, labeled carbon dioxide, labeled organic acids, and labeled amino acids in animals exposed to stable isotopes either from early development on nematode growth media agar plates or beginning as young adults while exposed to various pharmacologic treatments in liquid culture. Free amino acids are quantified by high performance liquid chromatography (HPLC) in whole worm aliquots extracted in 4% perchloric acid. Universally labeled 13C-glucose or 1,6-13C2-glucose is utilized as the stable isotopic precursor whose labeled carbon is traced by mass spectrometry in carbon dioxide (both atmospheric and dissolved) as well as in metabolites indicative of flux through glycolysis, pyruvate metabolism, and the tricarboxylic acid cycle. Representative results are included to demonstrate effects of isotope exposure time, various bacterial clearing protocols, and alternative worm disruption methods in wild-type nematodes, as well as the relative extent of isotopic incorporation in mitochondrial complex III mutant worms (isp-1(qm150)) relative to wild-type worms. Application of stable isotopic profiling in living nematodes provides a novel capacity to investigate at the whole animal level real-time metabolic alterations that are caused by individual genetic disorders and/or pharmacologic therapies.

Stable Isotopic Profiling of Intermediary Metabolic Flux in Developing and Adult Stage Caenorhabditis elegans

Stable isotopic profiling by gas chromatography mass spectrometric analysis of intermediary metabolic flux is described in the nematode, Caenorhabditis elegans. Methods are detailed for assessing isotopic enrichment in carbon dioxide, organic acids, and amino acids following isotope exposure either during development on agar plates or during adulthood in liquid culture.

Stable isotopic profiling has long permitted sensitive investigations of the metabolic consequences of genetic mutations and/or pharmacologic therapies in ...

Expression and Purification of the Cystic Fibrosis Transmembrane Conductance Regulator Protein in Saccharomyces cerevisiae

The cystic fibrosis transmembrane conductance regulator (CFTR) is a chloride channel, that when mutated, can give rise to cystic fibrosis in humans.There is therefore considerable interest in this protein, but efforts to study its structure and activity have been hampered by the difficulty of expressing and purifying sufficient amounts of the protein1-3. Like many 'difficult' eukaryotic membrane proteins, expression in a fast-growing organism is desirable, but challenging, and in the yeast S. cerevisiae, so far low amounts were obtained and rapid degradation of the recombinant protein was observed 4-9. Proteins involved in the processing of recombinant CFTR in yeast have been described6-9 .In this report we describe a methodology for expression of CFTR in yeast and its purification in significant amounts. The protocol describes how the earlier proteolysis problems can be overcome and how expression levels of CFTR can be greatly improved by modifying the cell growth conditions and by controlling the induction conditions, in particular the time period prior to cell harvesting. The reagants associated with this protocol (murine CFTR-expressing yeast cells or yeast plasmids) will be distributed via the US Cystic Fibrosis Foundation, which has sponsored the research. An article describing the design and synthesis of the CFTR construct employed in this report will be published separately (Urbatsch, I.; Thibodeau, P. et al., unpublished). In this article we will explain our method beginning with the transformation of the yeast cells with the CFTR construct - containing yeast plasmid (Fig. 1). The construct has a green fluorescent protein (GFP) sequence fused to CFTR at its C-terminus and follows the system developed by Drew et al. (2008)10. The GFP allows the expression and purification of CFTR to be followed relatively easily. The JoVE visualized protocol finishes after the preparation of microsomes from the yeast cells, although we include some suggestions for purification of the protein from the microsomes. Readers may wish to add their own modifications to the microsome purification procedure, dependent on the final experiments to be carried out with the protein and the local equipment available to them. The yeast-expressed CFTR protein can be partially purified using metal ion affinity chromatography, using an intrinsic polyhistidine purification tag. Subsequent size-exclusion chromatography yields a protein that appears to be &gt;90% pure, as judged by SDS-PAGE and Coomassie-staining of the gel.

Expression and Purification of the Cystic Fibrosis Transmembrane Conductance Regulator Protein in Saccharomyces cerevisiae

Attempts to express the cystic fibrosis transmembrane conductance regulator (CFTR) in Saccharomyces cerevisiae have, until now, yielded relatively low amounts of protein. This protocol and the associated reagents distributed via the Cystic Fibrosis Foundation should allow the preparation of milligram amounts of this 'difficult' eukaryotic membrane protein.

The  cystic fibrosis transmembrane conductance regulator (CFTR) is a chloride  channel, that when mutated, can give rise to cystic fibrosis in  ...

An Innovative Method for Exosome Quantification and Size Measurement

Although the biological importance of exosomes has recently gained an increasing amount of scientific and clinical attention, much is still unknown about their complex pathways, their bioavailability and their diverse functions in health and disease. Current work focuses on the presence and the behavior of exosomes (in vitro as well as in vivo) in the context of different human disorders, especially in the fields of oncology, gynecology and cardiology.
Unfortunately, neither a consensus regarding a gold standard for exosome isolation exists, nor is there an agreement on such a method for their quantitative analysis. As there are many methods for the purification of exosomes and also many possibilities for their quantitative and qualitative analysis, it is difficult to determine a combination of methods for the ideal approach. 
Here, we demonstrate nanoparticle tracking analysis (NTA), a semi-automated method for the characterization of exosomes after isolation from human plasma by ultracentrifugation. The presented results show that this approach for isolation, as well as the determination of the average number and size of exosomes, delivers reproducible and valid data, as confirmed by other methods, such as scanning electron microscopy (SEM).

A method for isolation of exosomes from whole blood and further analysis by nanoparticle tracking using a semi-automatic instrument is presented in this article. The presented technology provides an extremely sensitive method for visualizing and analyzing particles in liquid suspension.

Although the biological importance of exosomes has recently gained an increasing amount of scientific and clinical attention, much is still unknown about ...

Purification of the Cystic Fibrosis Transmembrane Conductance Regulator Protein Expressed in Saccharomyces cerevisiae

Defects in the cystic fibrosis transmembrane conductance regulator (CFTR) protein cause cystic fibrosis (CF), an autosomal recessive disease that currently limits the average life expectancy of sufferers to &#60;40 years of age. The development of novel drug molecules to restore the activity of CFTR is an important goal in the treatment CF, and the isolation of functionally active CFTR is a useful step towards achieving this goal.
We describe two methods for the purification of CFTR from a eukaryotic heterologous expression system, S. cerevisiae. Like prokaryotic systems, S. cerevisiae can be rapidly grown in the lab at low cost, but can also traffic and posttranslationally modify large membrane proteins. The selection of detergents for solubilization and purification is a critical step in the purification of any membrane protein. Having screened for the solubility of CFTR in several detergents, we have chosen two contrasting detergents for use in the purification that allow the final CFTR preparation to be tailored to the subsequently planned experiments.
In this method, we provide comparison of the purification of CFTR in dodecyl-&#946;-D-maltoside (DDM) and 1-tetradecanoyl-sn-glycero-3-phospho-(1&#39;-rac-glycerol) (LPG-14). Protein purified in DDM by this method shows ATPase activity in functional assays. Protein purified in LPG-14 shows high purity and yield, can be employed to study post-translational modifications, and can be used for structural methods such as small-angle X-ray scattering and electron microscopy. However it displays significantly lower ATPase activity.

Purification of the Cystic Fibrosis Transmembrane Conductance Regulator Protein Expressed in Saccharomyces cerevisiae

Heterologous expression and purification of the cystic fibrosis transmembrane conductance regulator (CFTR) are significant challenges and limiting factors in the development of drug therapies for cystic fibrosis. This protocol describes two methods for the isolation of milligram quantities of CFTR suitable for functional and structural studies.&#160;

Defects in the cystic fibrosis transmembrane conductance regulator (CFTR) protein cause cystic fibrosis (CF), an autosomal recessive disease that ...

A Standardized Method for the Analysis of Liver Sinusoidal Endothelial Cells and Their Fenestrations by Scanning Electron Microscopy

Liver sinusoidal endothelial cells are the gateway to the liver, their transcellular fenestrations allow the unimpeded transfer of small and dissolved substances from the blood into the liver parenchyma for metabolism and processing. Fenestrations are dynamic structures - both their size and/or number can be altered in response to various physiological states, drugs, and disease, making them an important target for modulation. An understanding of how LSEC morphology is influenced by various disease, toxic, and physiological states and how these changes impact on liver function requires accurate measurement of the size and number of fenestrations. In this paper, we describe scanning electron microscopy fixation and processing techniques used in our laboratory to ensure reproducible specimen preparation and accurate interpretation. The methods include perfusion fixation, secondary fixation and dehydration, preparation for the scanning electron microscope and analysis. Finally, we provide a step by step method for standardized image analysis which will benefit all researchers in the field.

The fenestrated liver sinusoidal endothelial cell is a biologically important filter system that is highly influenced by various diseases, toxins, and physiological states. These changes significantly impact on liver function. We describe methods for the standardisation of the measurement of the size and number of fenestrations in these cells.

Liver sinusoidal endothelial cells are the gateway to the liver, their transcellular fenestrations allow the unimpeded transfer of small and dissolved ...

Relating Stomatal Conductance to Leaf Functional Traits

Leaf functional traits are important because they reflect physiological functions, such as transpiration and carbon assimilation. In particular, morphological leaf traits have the potential to summarize plants strategies in terms of water use efficiency, growth pattern and nutrient use. The leaf economics spectrum (LES) is a recognized framework in functional plant ecology and reflects a gradient of increasing specific leaf area (SLA), leaf nitrogen, phosphorus and cation content, and decreasing leaf dry matter content (LDMC) and carbon nitrogen ratio (CN). The LES describes different strategies ranging from that of short-lived leaves with high photosynthetic capacity per leaf mass to long-lived leaves with low mass-based carbon assimilation rates. However, traits that are not included in the LES might provide additional information on the species&#39; physiology, such as those related to stomatal control. Protocols are presented for a wide range of leaf functional traits, including traits of the LES, but also traits that are independent of the LES. In particular, a new method is introduced that relates the plants&#8217; regulatory behavior in stomatal conductance to vapor pressure deficit. The resulting parameters of stomatal regulation can then be compared to the LES and other plant functional traits. The results show that functional leaf traits of the LES were also valid predictors for the parameters of stomatal regulation. For example, leaf carbon concentration was positively related to the vapor pressure deficit (vpd) at the point of inflection and the maximum of the conductance-vpd curve. However, traits that are not included in the LES added information in explaining parameters of stomatal control: the vpd at the point of inflection of the conductance-vpd curve was lower for species with higher stomatal density and higher stomatal index. Overall, stomata and vein traits were more powerful predictors for explaining stomatal regulation than traits used in the LES.

Unraveling how physiology and morphology are linked allows a deeper understanding of mechanistic functioning of plant leaves. We present both a procedure to derive parameters of stomatal regulation from stomatal conductance measurements and correlations with traditional functional leaf traits.

Leaf functional traits are important because they reflect physiological functions, such as transpiration and carbon assimilation. In particular, ...

Cardiac Pressure-Volume Loop Analysis Using Conductance Catheters in Mice

Cardiac pressure-volume loop analysis is the &#8220;gold-standard&#8221; in the assessment of load-dependent and load-independent measures of ventricular systolic and diastolic function. Measures of ventricular contractility and compliance are obtained through examination of cardiac response to changes in afterload and preload. These techniques were originally developed nearly three decades ago to measure cardiac function in large mammals and humans. The application of these analyses to small mammals, such as mice, has been accomplished through the optimization of microsurgical techniques and creation of conductance catheters. Conductance catheters allow for estimation of the blood pool by exploiting the relationship between electrical conductance and volume. When properly performed, these techniques allow for testing of cardiac function in genetic mutant mouse models or in drug treatment studies. The accuracy and precision of these studies are dependent on careful attention to the calibration of instruments, systematic conduct of hemodynamic measurements and data analyses. We will review the methods of conducting pressure-volume loop experiments using a conductance catheter in mice.

Cardiac pressure-volume loop analysis is the most comprehensive way to measure cardiac function in the intact heart. We describe a technique to perform and analyze cardiac pressure volume loops, using conductance catheters.

Cardiac pressure-volume loop analysis is the &#8220;gold-standard&#8221; in the assessment of load-dependent and load-independent measures of ventricular ...

Simple and Effective Administration and Visualization of Microparticles in the Circulatory System of Small Fishes Using Kidney Injection

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, implantation of transgenic cells and tiny optical sensors. However, intravenous microinjections into small animals, which are mostly used in biological and veterinary laboratories, are very difficult and require trained personnel. Herein, we demonstrate a robust and efficient method for the introduction of microparticles into the circulatory system of adult zebrafish (Danio rerio) by injection into the fish kidney. To visualize the introduced microparticles in the vasculature, we propose a simple intravital imaging technique in fish gills. In vivo monitoring of the zebrafish blood pH was accomplished using an injected microencapsulated fluorescent probe, SNARF-1, to demonstrate one of the possible applications of the described technique. This article provides a detailed description of the encapsulation of pH-sensitive dye and demonstrates the principles of the quick injection and visualization of the obtained microcapsules for in vivo recording of the fluorescent signal. The proposed method of injection is characterized by a low mortality rate (0-20%) and high efficiency (70-90% success), and it is easy to institute using commonly available equipment. All described procedures can be performed on other small fish species, such as guppies and medaka.

This article demonstrates the principles of a quick, minimally invasive injection of fluorescent microparticles into the circulatory system of small fishes and the in vivo visualization of the microparticles in fish blood.

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, ...

Measurement of Energy Metabolism in Explanted Retinal Tissue Using Extracellular Flux Analysis

High acuity vision is a heavily energy-consuming process, and the retina has developed several unique adaptations to precisely meet such demands while maintaining transparency of the visual axis. Perturbations to this delicate balance cause blinding illnesses, such as diabetic retinopathy. Therefore, the understanding of energy metabolism changes in the retina during disease is imperative to the development of rational therapies for various causes of vison loss. The recent advent of commercially-available extracellular flux analyzers has made the study of retinal energy metabolism more accessible. This protocol describes the use of such an analyzer to measure contributions to retinal energy supply through its two principle arms - oxidative phosphorylation and glycolysis - by quantifying changes in oxygen consumption rates (OCR) and extracellular acidification rates (ECAR) as proxies for these pathways. This technique is readily performed in explanted retinal tissue, facilitating assessment of responses to multiple pharmacologic agents in a single experiment. Metabolic signatures in retinas from animals lacking rod photoreceptor signaling are compared to wild-type controls using this method. A major limitation in this technique is the lack of ability to discriminate between light-adapted and dark-adapted energy utilization, an important physiologic consideration in retinal tissue.

This technique describes real time recording of oxygen consumption and extracellular acidification rates in explanted mouse retinal tissues using an extracellular flux analyzer.

High acuity vision is a heavily energy-consuming process, and the retina has developed several unique adaptations to precisely meet such demands while ...

Measuring Diurnal Rhythms in Autophagic and Proteasomal Flux

Cells employ several methods for recycling unwanted proteins and other material, including lysosomal and non-lysosomal pathways. The main lysosome-dependent pathway is called autophagy, while the primary non-lysosomal method for protein catabolism is the ubiquitin-proteasome system. Recent studies in model organisms suggest that the activity of both autophagy and the ubiquitin-proteasome system is not constant across the day but instead varies according to a daily (circadian) rhythm. The ability to measure biological rhythms in protein turnover is important for understanding how cellular quality control is achieved and for understanding the dynamics of specific proteins of interest. Here we present a standardized protocol for quantifying autophagic and proteasomal flux in vivo that captures the circadian component of protein turnover. Our protocol includes details for mouse handling, tissue processing, fractionation, and autophagic flux quantification using mouse liver as the starting material.

We describe our protocol for measuring biological rhythms in protein catabolism via autophagy and the proteasome in mouse liver.