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General
This study was carried out in the Lijiang Forest Ecosystem Research Station, Yunnan Province, China during the period 13 July to 3 August 2019. This field station, which is operated by the Kunming Institute of Botany, Chinese Academy of Sciences, is located to the north of Lijiang on Yulong Snow Mountain at an elevation of 3200 m. Canopy vegetation is dominated by Pinus yunnanensis and Quercus variabilis, and Rhododendron decorum is a highly conspicuous understory shrub species, when in flower. Flowers occur in inflorescences with each plant typically having many inflorescences.
We chose, numbered and bagged one inflorescence on each of 25 plants of Rhododendron decorum, followed the state of individual flowers, and sampled nectar according to a protocol explained below. We selected plants, as encountered, that were flowering and within about 10 m of our walking path, which was along a road and foot track near the field station. We selected inflorescences, one per plant, with at least five unopened buds, and marked five of these buds with small lengths of differently coloured plastic drinking straws22. The different colours enabled us to distinguish flowers during nectar sampling and subsequent measurements of flower colour. All marked flowers were then checked daily to record flower state as bud, beginning to open, open-non-abscised, and open-abscised. Flowers were considered buds if there was no sign of petals unfolding, beginning to open if petals had begun to unfold, and open if petals had unfolded completely. Abscised flowers were clearly indicated by separation between the base of the petals and the rest of a flower, which was along a distinct abscission line (Fig. 1c). Inflorescences were bagged, using green mesh organza bags, to prevent any flower visitation and nectar removal. This species is self-incompatible19, so no pollination occurred.
We carried out two experiments, involving a total of 25 plants. One (Experiment A) involved 10 plants (numbered A-1 to A-10) and was carried out between 13 and 23 July 2019. Experiment B involved 15 plants (numbered B-1 to B-15) and was carried out between 24 July and 3 August 2019. Experiment B was carried out to increase sample sizes for flowers of all ages, and to provide information for relatively young flowers that was not provided by Experiment A (explained further below). Plants were numbered as encountered.
Collection of inflorescences
Inflorescences from Experiment A and Experiment B were collected for sampling of nectar according to the following protocol.
For all inflorescences in Experiment A (i.e., 10 inflorescences) and all in Experiment B, except numbers B-3, 6, 9, 12 & 15 (i.e., 10 inflorescences), each inflorescence was removed from its plant on the first day that abscission of any marked flower was observed. If any marked flower was observed to have abscised, its inflorescence was removed from its plant by breaking its subtending stem and taken to a nearby sheltered ‘nectar sampling station’ where nectar measurements were made. This occurred for flowers between 3 and 9 days of age, counting the first day that a flower was either open or beginning to open as age 1. In a small number of cases, flower abscission occurred when marked flowers were gently touched just prior to nectar sampling. Such flowers were also considered to have abscised.
In addition, five inflorescences from Experiment B (i.e., numbers B-3, 6, 9, 12 & 15) were similarly collected when they were four days old, regardless of whether any flowers had abscised. This provided nectar measurements for relatively young flowers (i.e., ages 1 to 4 days).
Nectar sampling
For collected inflorescences, almost all the marked flowers were open, and we sampled accumulated nectar in each marked and open flower as follows. Nectar was removed using micro-capillary tubes (Hirschmann microcapillary pipettes; 5 µl in Experiment A; 10 µl in Experiment B; both 32 mm long), with volume measured on the basis of nectar length along tube and subsequently converted to µl. When about 0.5 µl of nectar was obtained, this was expelled to a hand-held refractometer (i.e., Bellingham & Stanley, 0 to 50% brix, adjusted for small volumes) for measurement of sugar concentration as % wt/wt sucrose equivalents. These measurements were adjusted for ambient temperature (see Supplementary Information) using a formula developed from information supplied by the manufacturers of the refractometers we use23 and converted to wt/vol using the following formula24: Y = 0.00226 + 0.00937X + 0.0000585X2 where Y is sugar mass per unit volume (mg/µl) and X is % concentration wt/wt. The amount of sugar for a flower (in mg) was then calculated by multiplying nectar volume (µl) by sugar mass per unit volume (mg/ µl).
Nectar was sampled, for both abscised and non-abscised flowers, from where it accumulates after secretion (Fig. 1c). Nectar was sampled for non-abscised flowers from the base of the corolla between the ring of about 10–15 nectaries, around the base of the ovary, and adjacent flower petals. For flowers that had abscised, nectar was separately sampled from both the ring of nectaries and the inside lowest 5 mm of the flower petals, where some nectar becomes attached.
Some flowers were judged to have been affected by rain and their nectar concentration measurements were excluded from analyses. There were periods of rain during our study and occasionally nectar concentration readings of lower than 1.5% wt/wt were obtained (n = 6), and the nectar assumed to have been diluted by rainwater. These records were excluded from analyses. Fortunately, most flowers pointed downwards and were thus not affected by rain.
Flower colour and pigment
Flower colours were measured by means of a modified Panasonic GH-1 camera. The low-pass filter of the camera had been removed in order to increase the sensitivity for ultraviolet light. The camera body was combined to an Ultra-Achromatic-Takumar 1:4.5/85 lens made of fused quartz that transmits UV light. Since the modified camera is sensitive to ultraviolet and infrared light, a UV-/IR-Cut filter transmitting light between 400 nm and 700 only nm was used to capture a normal reference picture. In addition, a UV-picture was captured from the identical position using a Baader UV-filter that transmits near ultraviolet light only. A white Teflon disc reflecting equal amounts of light in a range of wavelength from 300 to 700 nm was used for manual white balance before taking pictures. Using Image J both pictures were split into the RGB color channels, and then a false color photo was merged using the green channel of the color picture as red, the blue channel of the color picture as green, and the blue channel of the UV picture as blue (see Supplementary Information). For more details see article by Verhoeven et al.25. Using IrfanView image’s histogram a uniform non-decomposed area (number of pixels  > 10,000) of the adaxial corolla on the false color picture was selected. The average intensity for the red, green and blue channel of the false color photos with values between 0 and 255 was used for color evaluation. Abscised and non-abscised flowers were photographed together enabling direct comparison of the colours of the flowers.
Pigment content was deduced from the sum of the values of the red, green and blue channel of the false color photos. Since abscised and non-abscised flowers both appear white to the human eye, the possible change in the content of a UV-absorbing pigment was checked by comparing the value for the blue channel in relation to the sum for the values of the green and red channels.
Recordings of the spectral reflectance were done with an abscised and an open, non-abscised flower from each of five inflorescences. Reflectance measurements were performed with an USB2000 + spectrophotometer (Ocean Optics) and illumination was provided by a DH-2000-BAL light-source (Ocean Optics), both connected via a coaxial fibre cable. All measurements were taken in an angle of 90° to the measuring spot with a pellet of barium sulphate used as white standard and a black piece of plastic used as black standard.
Analyses
We used the General Linear Model approach to determine relationships for all flowers between nectar attributes (i.e., volume—µl, concentration—wt/vol, sugar weight—µg) as dependent variables and flower age, whether abscised, experiment (i.e., A vs. B), and Plant ID as independent variables. We also treated Plant ID as an independent categorical variable, but nested within experiment.
We used ANOVA to evaluate relationships between reflectance intensity and whether flower abscised, across different false colours, with Kolmogorov–Smirnov test for normality and Tukey post-hoc comparisons between means. We took log intensity as the dependent variable in order to meet the normality assumption.
We compared spectral reflectance for abscised and open, non-abscised flowers on the basis of the average reflectance across all wavelengths. Here we assumed that the 1140 reflectance values for each flower could be combined into a single average measure and that this average measure adequately represented each flower. We compared the two groups of flowers with a Kolmogorov–Smirnov Two Sample Test.
All analyses were carried out using the software SYSTAT26. More

Soils and their microbial communities
The soil properties shown in Table 1 indicate variation in soil texture across the Broadbalk field, with less clay in the N0 and FYM plots, situated on the north side of Broadbalk field compared to N6 and woodland towards the south side. The soil pH ranged from 7.1 to 8.2, lowest in the mineral-nitrogen fertilized soil N6 and highest in the N0 soil that received no N fertilizer. The bulk density of woodland soil is much lower and the % SOC much higher compared to other soils; the FYM soil has lower bulk density and higher % SOC than the other arable soils. The ratio of SOC:total N was approximately 10:1 in the arable soils and 13:1 in the woodland soil.
The community structure of bacteria and archaea revealed by 16S rRNA amplicon sequencing of metagenomic DNA extracted from the soil samples, at collection from the field, shows significant differences, and distinct separation on a NMDS plot (Fig. 1). Of the 14 phyla (sub-phyla for the Proteobacteria) comprising  > 0.1% of the community in at least one of the soils, only the δ-Proteobacteria did not show significantly different (P ≤ 0.05) mean abundance in at least one soil, according to ANOVA (Fig. 1). For example, the woodland soil has more α-Proteobacteria and Verrucomicrobia but fewer Thaumarchaeota (archaea) and β-Proteobacteria than the other treatments. Both the FYM and woodland soil have more γ-Proteobacteria and fewer Gemmatimonadetes; the FYM soil has more Firmicutes than the other soils (Fig. 1).
Figure 1

Relative mean abundance of prokaryotic phyla/sub-phyla in soils of origin on collection from the field. Phyla with at least 0.1% of the total community present in at least one soil treatment are included. Proteobacteria sub-phyla: a = alpha, b = beta, d = delta, g = gamma; s.e.d. for each group is shown; letters indicate mean significantly different means within each group (P = 0.05, according to Tukey’s post-hoc test on ANOVA). Insert top right shows NMDS plot of OTU for prokaryotic communities – PERMANOVA F = 9.477, P (same) = 0.0001.

Full size image

16S rRNA and denitrification gene abundance
At the end of the experiment, DNA was extracted and amplified from all samples but sufficient RNA for further analysis was obtained only from the FYM and woodland soils which contained more organic matter and larger microbial communities. ANOVA comparing the abundance for each set of genes and transcripts measured using qPCR showed that the soil of origin had a significant influence in all cases (Table 2). However, other factors (presence/absence of wheat and addition or not of N-fertilizer) and interactions between them were not significant, except for nosZI which was significantly influenced by the plant. Bacterial abundance indicated by 16S rRNA gene copy number was 2 × 109 g−1 soil in the N0 and N6 soils and significantly higher in the FYM and woodland soil, 5 × 109 and 7 × 109 copies g−1, respectively (Fig. 2). This pattern of relative abundance was seen for nirK (7 × 108–4 × 109 copies), nosZI (5 × 107–2 × 108 copies) and nosZII (4 × 106–1 × 107 copies g−1 soil). The exception was nirS where N0, N6 and woodland soil had similar gene abundance (1 × 107 copies g−1 soil) and FYM significantly more with 4 × 107 copies g−1soil (Fig. 2). The ratio nirK:nirS gene copies in the woodland soil was 300:1, significantly more than the mean of 55:1 in the arable soils (F3,32 = 102.63, P  More

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Site description and sampling
This experimental site was established in July 2009 at the Kessler Atmospheric and Ecological Field Station (KAEFS) in the US Great Plains in McClain County, Oklahoma (34̊ 59ʹN, 97̊ 31ʹW)14,48. Experimental design and site description were described in detail previously25. Briefly, Ambrosia trifida, Solanum carolinense and Euphorbia dentate belonging to C3 forbs, and Tridens flavus, Sporobolus compositus and Sorghum halapense belonging to C4 grasses are dominant in the site25,48. Annual mean temperature is 16.3 °C and annual precipitation is 914 mm, based on Oklahoma Climatological Survey data from 1948 to 1999. The soil type of this site is Port–Pulaski–Keokuk complex with 51% of sand, 35% of silt and 13% of clay, which is a well-drained soil that is formed in loamy sediment on flood plains. The soil has a high available water holding capacity (37%), neutral pH and 1.2 g cm−3 bulk density with 1.9% total organic matter and 0.1% total nitrogen (N)25,48. Four blocks were used in the field site experiment, in which warming is a primary factor. Two levels of warming (ambient and +3 °C) were set for four pairs of 2.5 m × 1.75 m plots by utilizing a real or dummy infrared radiator (Kalglo Electronics, Bethlehem, PA, USA). In the warmed plots, a real infrared radiator was suspended 1.5 m above the ground, and the dummy infrared radiator was suspended to simulate a shading effect of the device in the control plots.
In this study, eight surface (0–15 cm) soil samples, four from the warmed and four from the control plots, were collected annually at approximately the date of peak plant biomass (September or October) from 2010 to 2016. Three soil cores (2.5 cm diameter × 15 cm depth) were taken by using a soil sampler tube in each plot and composited to have enough samples for soil chemistry, microbiology and molecular biology analyses. A total of 56 soil samples were analyzed in this study.
Environmental and soil chemical measurements
Precipitation data were obtained from the Oklahoma Mesonet Station (Washington Station)48 located 200 m away from our experiment site, and 12-month version of the standardized precipitation-evapotranspiration index (SPEI-12) was used as annual drought index49. Air temperature, soil temperature and volumetric soil water content were described in detail previously25. Specifically, air temperature and soil temperature at the depth of 7.5 cm in the center of each field plot were measured by using Constantan-copper thermocouples wired to a Campbell Scientific CR10x data logger (Campbell Scientific, UT, USA). A portable time domain reflectometer (Soil Moisture Equipment Corp.) was used to measure soil moisture from the soil surface to a 15-cm depth once or twice a month. Three measurements of soil moisture were performed in each plot and the average of three technical replicates were used in further analyses.
All soil samples were analyzed to determine soil total organic carbon (TOC), total nitrogen (TN), soil nitrate (NO3−) and ammonia (NH4+) by the Soil, Water, and Forage Analytical Laboratory at Oklahoma State University (Stillwater, OK, USA). Soil pH was measured using a pH meter with a calibrated combined glass electrode50.
Aboveground plant communities
Aboveground plant community investigations were annually conducted at peak biomass (usually September)48,51. Aboveground plant biomass, separated into C3 and C4 species, was indirectly estimated by a modified pin-touch method48,51. Detailed description of biomass estimation is provided by Sherry et al.52. A pin frame used in this study is 1 m long and have 10 pins 10 cm apart at 30° from vertical. Pins with a 0.75 m length were raised within the frame to count hits up to 1 m high (hits over 1 m are negligible at this site). The pin frame was placed in the center of each plot to record the contact numbers of the pins separately with C3 and C4 plants (e.g., leaves and stems). The contact numbers of C3 and C4 plants were then used to estimate plant biomass using calibration equations derived from calibration plots, which were located near the experimental plots. Biomass in the calibration plots was clipped at a height of 10 cm above the ground at approximately the date of peak plant biomass (September or October). All of the species in plant community within each plot were identified to estimate species richness. Clipped plant materials were oven-dried and then correlated with the total contact number. C3 and C4 plant biomasses were estimated by using the calibration equation of contact number and plant biomass. All of the species within each plot were identified to estimate species richness of plants.
Ecosystem C fluxes and soil respiration
Ecosystem C fluxes and soil respirations were measured once or twice a month between 10:00 and 15:00 (local time) from January 2010 to December 2016 by following previous methods14,48. One square aluminum frame (0.5 m × 0.5 m) was inserted in the soil at 2 cm depth in each plot to provide a flat base between the soil surface and the CO2 sampling chamber. NEE and ecosystem respiration (ER) were measured using LI-6400 portable photosynthesis system (LI-COR). Gross primary productivity (GPP) was estimated as the difference between NEE and ER. Meanwhile, soil surface respiration was monthly measured using a LI-8100A soil flux system attached to a soil CO2 flux chamber (LI-COR). Measurements were taken above a PVC collar (80 cm2 in area and 5 cm in depth) and a PVC tube (80 cm2 in area and 70 cm in depth) in each plot. The PVC tube was permanently fixed on the ground to cut off old plant roots and prevent new roots from growing inside the tube. Any aboveground parts of living plants were removed from the PVC tubes and collars before each measurement. The CO2 efflux measured above the PVC tubes represented heterotrophic respiration (Rh) from soil microbes, while that measured above the PVC collars represented soil total respiration (Rt) including heterotrophic and autotrophic respiration (Rh and Ra) from soil microbes and plant root, respectively.
Soil decomposition rate
Weighted cellulose filter paper (Whatman CAT No. 1442-090) was placed into fiberglass mesh bags and placed vertically at 0–10 cm soil depth in each plot in March 2016. All of decomposition bags were collected back in September 2016, rinsed and dried at 60 °C for weighing. The percentage of mass loss was calculated to represent soil decomposition rate.
Molecular analyses of soil samples
The C substrate utilization patterns of soil microbial communities in 2016 were analyzed by BIOLOG EcoPlateTM (BIOLOG). The BIOLOG EcoPlateTM contains 31 of the most useful labile carbon sources for soil community analysis, which are repeated three times in each plate. In this study, the plates with diluted soil supernatant (0.5 g soil with 45 mL 0.85% NaCl) were incubated in a BIOLOG OmniLog PM System at 25 °C for 4.5 days. The color change of each well was shown as absorbance curve. The net area under the absorbance versus time curve was calculated to represent physiological activity of various C sources53. The average value from three replicates was used for analyses in this study.
Soil total DNA was extracted from 1.5 g soil by freeze-grinding and SDS-based lysis54, and purified with a MoBio PowerSoil DNA isolation kit (MoBio Laboratories)25. Then, 10 ng DNA per sample were used for library construction and amplicon sequencing. Amplicons sequencing was performed with cautions in terms of experimental preparations and data analyses to ensure sequence representativeness and semi-quantitative nature55. The V4 region of bacterial and archaeal 16S rRNA genes were amplified with the primer set 515F (5ʹ-GTGCCAGCMGCCGCGGTAA-3′) and 806R (5ʹ-GGACTACHVGGGTWTCTAAT-3ʹ), and fungal ITSs between 5.8S and 28S rRNA genes were amplified with the primer set ITS7F (5ʹ-GTGARTCATCGARTCTTTG-3ʹ) and ITS4R (5ʹ-TCCTCCGCTTATTGATATGC-3ʹ). PCR products from different samples were sequenced on a MiSeq platform (Illumina, Inc.) using 2 × 250 pair-end sequencing kit. Raw sequences were submitted to our Galaxy sequence analysis pipeline (http://zhoulab5.rccc.ou.edu:8080) to further analyze according to the protocol in the pipeline25. Finally, OTUs were clustered by UPARSE56 at 97% identity for both 16S rRNA gene and ITS. All sequences were randomly resampled to 30,000 sequences for 16S rRNA gene and 10,000 sequences for ITS per sample. Representative sequences of OTUs were annotated taxonomically by the Ribosomal Database Project (RDP) Classifier with 50% confidence estimates.
GeoChip 5.0 M, a functional gene array57, was used for all 56 samples from 2010 to 2016. GeoChip hybridization, scanning and data processing were performed in the Institute for Environmental Genomics, University of Oklahoma57,58. Specifically, 800 ng of purified soil DNA of each sample was mixed with 5.5 µl random primers (Life Technologies, random hexamers, 3 µg/µl), diluted with nuclease-free water to 35 µl, heated to 99 °C for 5 min, and placed on ice immediately. The labeling master mix (15 µl), including 0.5 µl of Cy-3 dUTP (25 nM; GE Healthcare), 2.5 µl of dNTP (2.5 mM dTTP, 5 mM dAGC-TP), 1 µl of Klenow (imer; San Diego, CA; 40 U ml−1), 5 µl Klenow buffer, and 2.5 µl of water, was added in the sample mixed solution. The samples were incubated at 37 °C for 6 h in a thermocycler, and then incubated at 95 °C for 3 min to inactivate the enzyme. Subsequently, samples were protected from the light as much as possible. Labeled DNA was cleaned using a QIAquick purification kit (Qiagen) according the manufacturer’s instructions and then dried thoroughly in a SpeedVac (45 °C, 45 min; ThermoSavant).
Labeled DNA was resuspended into 27.5 µl of DNase-free water, and then mixed completely with 99.4 µl of hybridization solution, containing 63.5 µl of formamide (10% final concentration), 2 × HI-RPM hybridization buffer, 12.7 µl of 10 × aCGH blocking agent, 0.05 μg/µl Cot-1 DNA, and 10 pM CORS58. The mixed solution was denatured at 95 °C for 3 min, and then incubated at 37 °C for 30 min. The DNA solution was centrifuged at 6000 × g for 1 min to collect liquid at the bottom of the tube. 110 µl of the solution was pipetted into the center of the well of the gasket slide. The array slide was placed on the gasket slide, sealed using a SureHyb chamber, hybridized at 67 °C for 24 h at 20 rpm in a hybridization oven. After hybridization, slides were washed in room temperature with Wash Buffer 1 (Agilent) and Wash Buffer 2 (Agilent).
The slides were imaged as a Multi-TIFF with a NimbleGen MS200 Microarray Scanner (Roche NimbleGen, Inc., Madison, WI, United States). The raw signals from NimbleGen were submitted to the Microarray Data Manager on our website (http://ieg.ou.edu/microarray), cleaned, normalized and analyzed using the data-analysis pipeline. Briefly, probe quality was assessed, and poor or low signal probes were removed. Probe spots with coefficient of variance (CV; probe signal SD/signal) >0.8 were removed. Then, the signal-to-noise ratio (SNR) was calculated. As suggested by Agilent, the average signal of Agilent’s negative control probes within each subarray was used as the background signal for the probes in that subarray instead of the local background typically used. The signal intensity for each spot was corrected by subtracting the background signal intensity. If the net difference was30%, aligned length >20 a.a., and e-value More