2.2 Acridine orange staining and anatomical characterization
Root segments (approximately 0.5 cm) were sampled in triplicate at 3 cm,
6 cm, 9 cm, and 12 cm from the taproot root tip 15 days after the
drought stress treatment. The root segments were stored in 75% ethanol
and phenotypically analyzed for the presence of RCS in the cortex using
acridine orange staining, as described by Henry and Deacon (1981).
Acridine orange staining is a viable method to assess RCS. Acridine
orange fluorescent nuclei were used for phenotyping the onset of RCS
within individual cortical layers. The disappearance of the cortex,
determined by the percentage of viable cells, was used to indicate RCS
manifestation at the root cross-sectional level. The acridine
orange-stained root segments were embedded in a gelatin capsule with
Tissue-Tek CRYO-OCT compound (Thermo Fisher Scientific, Waltham, MA,
USA) and snap-frozen in liquid nitrogen before storage at -20 °C.
Cross-sections (8 µm thick) of the embedded segments were cut using a
Freeze
Slicer (Leica CM1950) and
imaged
using a laser scanning confocal microscope (FV-1000, Olympus, Japan).
The main measurements using the
cross-sectional images included area determination and variable
counting. The following area measurements were obtained through pixel
counting: the total cross-section area and diameter, the stele area and
diameter, and the cortex area and diameter. The average cell size was
also calculated by pixel counting, and the cortical areas of acridine
orange fluorescent nuclei were measured. The number of acridine orange
fluorescent nuclei was counted in all cell layers of the 8 μm thick
sections. The count data included the number of cortical cells, cortical
cell files, and cortical lacunae area. The calculation formulas were as
follows (Schneider et al., 2017b; Schneider et al., 2017a; Schneider et
al., 2018; Schneider and Lynch, 2018):
\(\mathrm{Cortical\ area=Cross\ section\ area\ }\mathrm{\ }\mathrm{\text{Stele\ area}}\)(1)\(\mathrm{Cortical\ diameter=Cross\ section\ diameter\ }\mathrm{\ }\mathrm{\text{Stele\ diameter}}\)(2)\(\mathrm{Stele/whole\ ratio=Stele\ area/Cross\ section\ area\ }\) (3)\(\mathrm{Lacunae/cort}\mathrm{\text{ex}}\mathrm{\ ratio=Total\ lacunae\ area/Cort}\mathrm{\text{ex\ }}\mathrm{\text{area}}\ \)(4)\(\mathrm{Cortex/stele\ ratio=Cross\ section\ area/Stele\ area}\) (5)
RCS-free control samples were root
segments collected at 3 cm from the root tip (Schneider et al., 2017b).
The cortical senescence percentage
was calculated by comparing the cortical area (total cross-sectional
area
-
stele area) at 3 cm from the root tip with the corresponding area on the
sampled position (Schneider et al., 2017b).
2.3
The root metabolic enzyme
activity and root respiration
Root
segments
(approximately 0.5 cm) sampled at 3 cm, 6 cm, 9 cm, and 12 cm from the
taproot root tip 15 days after drought stress treatment were used to
measure root metabolic enzyme activity and root
respiration. The measurements were
conducted in triplicate.
Phosphofructokinase activity levels were determined using the method
described by Baldwin et al. (2007), and changes in absorbance were
monitored continuously at 340 nm.
Malate dehydrogenase activity levels were determined using the method
described by Childress and Somero (1979), whereby the NADH oxidation was
quantified at 340 nm.
Glucose-6-phosphate
dehydrogenase activity induced by the reduction of NADP+ to NADPH was
measured at 340 nm (Antonietta Ciardiello et al.,
1995).
Root respiration was measured for 10 minutes using a LI-840A
CO2/H2O gas analyzer (LI-COR, Inc.,
Lincoln, NE, USA), after which the root segment was dried and weighed
(Sun et al., 2021).
2.4 Endogenous hormone
content
Root segments were sampled as described in section 2.3.
Phytohormone,
gibberellin
(GA), zeatin riboside (ZR), indole-3-acetic acid (IAA), brassinolide
(BR) and abscisic acid (ABA) contents in the root segments were
determined using an Agilent 1260 High-Performance Liquid Chromatograph
with an Eclipse XDB-C18 column (250 mm × 4.6 mm, 5 mm, Agilent, CA,
United States).
Briefly, 2 g of fresh root segments were ground in 10 mL of pre-cooled
80% methanol in an ice bath. After centrifugation at 8,000 g for 10
minutes at 4°C, the supernatant was collected, and the residue was
resuspended in 8 mL pre-cooled 80% methanol, followed by another round
of centrifugation and supernatant collection. The aqueous phase was then
extracted three times with 20 mL of ethyl acetate. The sample was
evaporated and dried at 40°C under pressure, and the separation of the
five endogenous phytohormones was conducted via gradient elution (Zhang
et al., 2013).
2.5
Root system phenotyping
Root samples were obtained in triplicate at 0, 5, 10, and 15 days after
drought stress
treatment.
Digital images with a resolution of 600 dots per inch were obtained
using a scanner (EPSON Expression 10000XL). The root images were
analyzed using WinRHIZO (Regent Instruments, Inc., Quebec City, Canada)
and RootNav software (Christopher et al., 2013; Pound et al., 2013) to
obtain information on the root traits (Table S1) (Zhang et al., 2021).
Thereafter, the fresh weight of the roots and aboveground parts was
obtained by weighing, and the biomass was obtained by drying at 105 °C
for 30 minutes and then at 75 °C for 72 hours to constant weight.
2.6 Morphological
and physiological traitsof the aboveground parts
Aboveground parts were sampled as
described in section 2.5. Plant height was measured using a
straightedge, and leaf area was calculated using the length and width
factor method. Stem diameter was recorded with a vernier caliper.
The ratio of variable to maximum fluorescence (F v/F m) was
measured using a portable Chlorophyll Fluorescence monitoring system
(FMS-2, Hansatech, King’s Lynn, UK). The relative chlorophyll content
was measured using a SPAD chlorophyll meter (SPAD-502; Konica Minolta,
Tokyo, Japan) while avoiding the leaf veins.
The
leaf water potential was measured with a Model 600 plant pressure
chamber (PMS, USA), while stomatal size and density were measured using
a combination of nail polish and sticky tape. The relative water content
and leaf water saturation deficit were determined by weighing (Yang et
al., 2022). These measurements were based on the third functional leaf
and were conducted between 9:00 and
11:00.