Plant material and growth conditions
Six-month-old tamarack seedlings were obtained from the Bonnyville Forest Nursery Inc., AB and stored frozen over winter. In spring, the seedlings were transferred into 4.5 L pots (Nursery Supplies, Tustin, CA) filled with commercial soil mixture (Sunshine LA4 Mix, Sun Grow Horticulture Canada Ltd, Spruce Grove, AB, Canada). The pots were placed in a greenhouse that was maintained at 22/18 °C day/night temperature, 18-h photoperiod, 65 ± 10% relative humidity. Natural light was supplemented with 400-W high pressure sodium lamps giving a minimum photosynthetic photon flux density (PPFD) of 350 μmol m-2 s-1 (Lumalux, GTE Sylvania, Drummondville, PQ, Canada). The seedlings were fertilized once a week with the 20:20:20 (N:P:K) commercial fertilizer applied at 0.25 g L-1.
After one month, a group of 20 seedlings was subjected to flooding for six months. The flooding treatment was applied by immersing the pots in mineral solution up to about 5 cm below the top edge. The mineral solution was changed once a week. The initial oxygen concentration of the solution after each weekly change was about 8 mg L-1, and declined over the course of the week to approximately 2–3 mg L-1.
A second group of 20 seedlings served as non-flooded control. The pots with control plants were placed on a greenhouse bench next to the flooded plants and the plants were regularly watered and fertilized as above. The experiment was repeated three times in three different years at the same times during the growing season (Study 1, 2, and 3).
Dry weights, gas exchange and leaf chlorophyll concentrations
Plant dry weights, gas exchange and leaf chlorophyll concentrations were measured after six months of flooding in Study 1. For dry weight determinations, roots were separated from shoots in six seedlings per treatment (n = 6) and their dry weights obtained after drying at 65 °C for 72 h.
Net photosynthesis (NP) and transpiration (E) rates were measured in the greenhouse between 10:00 and 11:30 using the uppermost branches of six seedlings (n = 6) per treatment after six months of the flooding treatment. At that time, the upper branches of flooded plants did not shown signs of needle chlorosis or necrosis, which were already apparent in the lower branches. For the gas exchange measurements, an infrared gas analyzer (LCA-4, Analytical Development Company Ltd., Hertfordshire, UK) was used with an auxiliary light source (1000 μmol m−2 s−1 PPFD). Needle surface areas were determined with the Sigma Scan 5.0 software following scanning (Jandel Scientific, San Rafael, CA).
Shoot water potential was measured with a Scholander pressure chamber in the upper lateral branches of the trees. The branches were attached from the stems into the pressure chamber and compressed air was applied. The water potential was determined as the pressure needed for the first drop of water to come out from the branch stem.
Needle chlorophyll concentrations were determined in six seedlings per treatment (n = 6) in the same needles as those used for the gas exchange measurements. The needles were freeze-dried and grinded. For chlorophyll determinations, 20 mg DW of tissue was extracted with 8 ml methanol. The methanol extracts were analyzed spectrophotometrically (Ultrospec III, Pharmacia LKB, Uppsala, Sweden) and the MacKinney’s equation  was used for the calculations of needle chlorophyll concentrations.
Root hydraulic conductance (KTOT, KIND), conductivity (LTOT, LIND) and xylem exudate concentrations of 1-hydroxypirene-3,6,8 – trisulfonic acid (PTS3)
Whole root system hydraulic conductance (KTOT) and conductivity (LTOT) were determined after six months of flooding treatment in six flooded and six non-flooded plants (n = 6) in Study 1. The plants were measured at room temperature from 10:00 to 16:00. The shoots of seedlings were excised about 1.5 cm above the root collar without removing the roots from the soil. The roots were attached to the high pressure flow meter (HPFM, Dynamax Inc., Houston, TX) for KTOT measurements. Each root system was gradually pressurized to 0.5 MPa . After the measurements, root volumes were calculated using the volume displacement method  and LTOT was calculated.
For Study 2, root hydraulic conductance measurements were carried out with a HPFM on sets of individual roots (KIND) cut at the stem level (Figure 2) of six non-flooded plants and on six adventitious roots from flooded seedlings (n = 6). For the measurements, the roots were placed in a water bath at 10 °C and their KIND was determined at increasing temperatures from 10, 15, 20, 25, to 30 °C. The roots were held for 10 minutes after each temperature change before the measurement. Root hydraulic conductivity (LIND) was calculated based on the root volume calculated by the volume displacement method as explained above.
The activation energy (Ea, kcal mol-1) for root water transport was calculated from the Arrhenius plots of the natural logarithm of LIND values against the inverse of absolute temperatures .
The apoplastic tracer dye 1-hydroxypirene-3,6,8 – trisulfonic acid (PTS3) was used to examine relative changes in the apoplastic to cell-to-cell water flow ratio in the roots. PTS3 is a fluorescent dye that is not transported across the cell membranes and which have been often used to estimate changes in ratio of apoplastic measurements of cell-to-cell water transport in roots. For the measurements, we used the same sets of roots as in the measurements of LIND. The roots were placed in a Scholander pressure chamber in an aqueous solution of 0.02% (w:v) PTS3, and pressurized at 0.3 MPa for 10 min . The exudates were collected and PTS3 concentrations were determined using a Sequoia-Turner 450 fluorometer (Apple Scientific, Chesterland, OH, USA) with 405 nm excitation and 515 nm emission spectra .
Root anatomy and root starch content
Root tips of eight control-non flooded seedlings and eight distal root segments of adventitious and non-adventitious flooded seedlings were randomly selected after 6 months of flooding treatment (between 0.5- and 1-cm long in Study 1 and up to 5-cm long in Study 3). In Study 1, the root segments were fixed with FAA (70% ethanol, glacial acetic acid and formalin) overnight followed by 4% paraformaldehyde and 0.5% glutaraldehyde for 6 h. After fixation, the root segments were dehydrated in an ethanol series, placed in toluene and embedded in paraffin. Serial cross sections (6-8-μm thick) of paraffin-embedded roots were prepared with a microtome. The paraffin sections were cleared with toluene and 95% ethanol and stained with 0.1% Safranin O (in 95% ethanol (w:v) for 1 h follow by 0.1% Fast Green FCF in 95% ethanol (w:v) for 1 min. The sections were mounted on slides and examined under the light microscope.
The development of xylem and endodermal tissues were examined in Study 3. Root segments were obtained at different distances from the fresh root tips (0, 0.5, 1, 1.5, 2, and 3 cm). Fresh, free-hand sections were prepared with a razor blade and stained with Sudan IV or Berberine . The sections were examined with the light and fluorescent microscope (Leica DMRXA Upright Microscope) using a green light filter I3 at 450–490 nm excitation and 510 nm emission.
Starch content and distribution was examined in fresh, free-hand sections taken up to 1 cm from the root tip of non-flooded roots and adventitious roots from flooded plants. The sections were stained with IKI and viewed under the light microscope.
Inmunolocalization of root aquaporins
For in situ immunolocalization of aquaporins, fresh free-hand sections were prepared with a razor blade from the roots of six non-flooded and flooded seedlings. The sections were taken from 0.5 to 1 cm from the root tip. They were incubated with the antibodies raised against PIP1 and PIP2 aquaporins . The anti-PIP1 antibody (R-4445) was raised against an amino peptide from ZmPIP1;5 and recognizes all maize PIP1s except PIP1;6 . The anti-PIP2 antibody (R-2493) was raised against an amino peptide from ZmPIP2;4 and was shown to recognize all maize PIP2s (Dr. F. Chaumont personal communication). Following incubation in blocking solution (5% BSA in phosphate buffer solution) and in the anti-PIP1, PIP2, and preimmune antibodies (1:1000 dilution by volume), the sections were incubated in the fluorescein-coupled goat anti-rabbit IgG antibody (Sigma-Aldrich Canada Ltd., ON, Canada). The slides were examined under the fluorescence microscope (Leica DMRXA Upright Microscope) with a green light filter I3 at 450–490 nm excitation and 510 nm emission.
To determine the specificity of polyclonal antibodies, sodium dodecyl sulfate-polyacrylamide gel electrophoresis (12% acrylamide resolving gel) was performed with proteins (30 μg) extracted from the roots of a two-year old tamarack (Larix laricina) and two-week old greenhouse-grown maize (Zea mays). Following electrophoresis, gels were electroblotted to PVDF membrane, blocked overnight in Tris-buffered saline containing 5% (w/v) bovine serum albumin and probed with primary polyclonal antibodies (1:500) raised against Zea mays PIP1 and PIP2 aquaporins that were used for the immunolocalization . Secondary antibody (1:10,000) raised in goat against rabbit IgG and conjugated to alkaline phosphatase (Sigma-Aldrich) was used to detect immunoreactive bands.
The data were analyzed using SAS (version 9.1, SAS Institute Inc.; Cary, NC, USA). T-test was used to determine significant differences between treatment means of water relation and physiological parameters at α = 0.05. Analysis of variance (ANOVA) with the Mixed Procedure of SAS was used to determine differences in LIND of individual roots at different temperatures at α = 0.05.