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Oecologia

, Volume 187, Issue 4, pp 1053–1075 | Cite as

Distinctions in heterotrophic and autotrophic-based metabolism as recorded in the hydrogen and carbon isotope ratios of normal alkanes

  • Brett J. Tipple
  • James R. Ehleringer
Special Topic

Abstract

The hydrogen isotope values of n-alkanes (δ2Hn-alkane) reflect a plant’s water source and water relations, while the carbon isotope values (δ13Cn-alkane) relate to a plant’s carbon metabolism and response to environmental conditions. However, the isotopic dynamics of the transition from heterotrophic to autotrophic metabolism during foliar development on δ2Hn-alkane and δ13Cn-alkane remain unclear. Here, we monitored δ2Hn-alkane and δ13Cn-alkane across a growing season from Betula occidentalis, Populus angustifolia, and Acer negundo. In addition, we compiled δ2H values of atmospheric vapor, leaf water, xylem water, and stream water as well as δ13C values of bulk leaf tissue (δ13Cbulk). We found δ2Hn-alkane and δ13Cn-alkane varied with leaf development and indicated that the majority of wax development occurred during the initial growing season. The patterns in δ2Hn-alkane were broadly consistent between species and with previous studies; however, each species had a unique final δ2Hn-alkane value. The δ13Cbulk for all species demonstrated a characteristic 13C-enrichment during the initial growing season, followed by 13C-depletion, while δ13Cn-alkane did not exhibit a consistent trend between the species. These δ13C data suggested a decoupling of the isotopic inputs between n-alkanes and photosynthetic leaf tissue. When coupled with δ2Hn-alkane, these data suggested that the precursor compounds utilized in initial production of n-alkanes might be variable and possibly indicated that the stored precursors used for initial leaf tissue and wax production originated from different sources. Nonetheless, these data indicated that the isotopic signatures of n-alkanes relate to a mixture of precursors, but only during a distinct period of leaf ontogeny.

Keywords

Leaf wax Cuticle Stable isotope Compound-specific isotope analysis Ecophysiology 

Introduction

Stable isotope analyses of plant tissues and compounds have become an indispensable tool to understand plant physiology, ecology, photosynthetic pathway, and growth conditions (e.g., Berke et al. 2015; Dawson and Ehleringer 1991, 1993; Ehleringer et al. 1986; Farquhar et al. 1989; Feakins and Sessions 2010a; Flanagan et al. 1991; Gessler et al. 2009; Helliker and Ehleringer 2002; Kahmen et al. 2011; Roden and Ehleringer 1999; Rundel et al. 1989; Sparks and Ehleringer 1997; Tipple and Pagani 2013; Zimmerman and Ehleringer 1990). Measurement of stable isotopes from historical and ancient plant materials, such as tree-ring cellulose, ancient soil organic matter, and sedimentary leaf wax lipids, is regularly used to reconstruct past ecologic and climatic conditions (Alstad et al. 2008; Cerling et al. 1994; Cook et al. 2010; Freeman and Colarusso 2001; Griffin and Anchukaitis 2014; Leavitt et al. 2011; McCarroll and Loader 2004; Smith et al. 2007; Treydte et al. 2009, among others). Recent advancements in compound-specific isotope ratio mass spectrometry and extraction methods have allowed for increased analysis throughput and development of higher resolution hydrogen (δ2H) and/or carbon (δ13C) isotope records of ancient plant ecology and hydrology from n-alkyl lipids (e.g., Berke et al. 2012; Diefendorf et al. 2015a; Douglas et al. 2015; Liddy et al. 2016; Tierney et al. 2013; Tipple et al. 2011; Uno et al. 2016). This has led to an intersection where ecologists, geologists, and climatologist share a common interest. As paleoclimatogists and geochemists have increased the applications of isotope analysis of leaf wax lipids to understand historical and ancient systems, plant ecologists, biochemists, and botanists have sought to better understand the biochemical and environmental controls on the δ2H or δ13C values of leaf waxes and other plant compounds (e.g., Gamarra and Kahmen 2015; Kahmen et al. 2013a, b; Ladd and Sachs 2017; Newberry et al. 2015; Oakes and Hren 2016; Sachse et al. 2010, 2015; Tipple et al. 2016; Zhang et al. 2017). The shared interest in these compounds has led to numerous synergistic collaborations between the biological, ecological, and geological communities (Diefendorf et al. 2015b; Lee et al. 2016; Sachse et al. 2012).

The δ2H plant leaf wax compounds, particularly the n-alkyl lipids, are powerful tools to understand and reconstruct a plant’s water environment (Dawson et al. 2002). The relationship between the δ2H of plant source water and n-alkyl lipids is well established, as source water is the ultimate source of hydrogen to all plant compounds (Sachse et al. 2012). The δ2H of n-alkyl lipids is also shaped by atmospheric exchange and leaf water 2H-enrichment from evaporation from the leaf (Kahmen et al. 2013b; Tipple et al. 2015) as well as the biosynthetic fractionations that occur during the formation of the lipids (Schmidt et al. 2003), all of which add to the information encoded in the δ2H values of n-alkyl lipids. The n-alkyl lipids (e.g., n-alkanes, fatty acids) are synthesized through the acetogenic pathway (Zhou et al. 2010). 3-phosphoglyeric acid, the precursor molecule for the acetogenic pathway and the intermediate, pyruvate, originating from the Calvin Cycle or a secondary metabolism. The acetogenic pathway results in a four-carbon butyryl chain with seven hydrogen atoms that through fatty acid synthesis is elongated to form a C16 or C18 fatty acid with the addition of two-carbon acetyl-CoA units (Zhou et al. 2010). On the initial butyryl chain, three hydrogen atoms originate from biosynthetic water, two from NADPH from the light reaction, and two from precursor acetyl-CoA from the chloroplast pyruvate (Cormier et al. 2018; Sachse et al. 2012; Schmidt et al. 2003; Zhou et al. 2010). Of the hydrogen atoms added during elongation within the chloroplast, 50% was derived from NADPH, 25% from the surrounding biosynthetic water, and the remaining 25% from chloroplast pyruvate (Cormier et al. 2018). These fatty acids are exported and further elongated in the endoplasmic reticulum, creating longer alkanoic acids and alkanes with hydrogen atoms derived from NADPH (50%), biosynthetic water (25%), and precursor pyruvate (25%) (Cormier et al. 2018). Several different metabolic pathways are used to produce NADPH and, thus, the relative importance of these pathways in the production of n-alkyl lipids will affect the δ2H value (Cormier et al. 2018). Nonetheless, the primary source of NADPH in higher plants is photosynthesis, as demonstrated by the strong linear relationship between the δ2H of source water and n-alkyl lipids (Sachse et al. 2012).

The δ13C of n-alkyl lipids is useful to understand modern and past plant physiology and ecophysiology (Diefendorf and Freimuth 2017). The δ13C values of plant material reflect photoassimilated CO2 and fractionations associated with carbon fixation. During photosynthesis, CO2 is fixed and converted to sugars and the δ13C values of photosynthate are most strongly affected by the discrimination of 13C by Rubisco—the enzyme that fixes CO2. The δ13C of photosynthate and its products is also a function of CO2 diffusion, leaf conductance, and the concentrations of intercellular and atmospheric CO2 (Farquhar et al. 1989). Also, photosynthetic pathway has a large control on the final δ13C of products. The C3 photosynthetic pathway is the most common and has the largest net fractionation between CO2 and the plant material. The C4 and Crassulacean acid metabolism (CAM) photosynthetic pathways spatially or temporally separate CO2 fixation from carbon assimilation, respectively. As a result, the δ13C of C4 and CAM plant materials are less negative than those of C3 plants. As with other plant tissues, the δ13C of lipids reflects and records the photosynthetic pathway of the plant. In higher plants, the initial steps of n-alkyl lipid biosynthesis occur in the chloroplast, with carbon atoms originating from chloroplast pyruvate (Zhou et al. 2010). The elongation phase occurs in the endoplasmic reticulum, with the carbon atoms added during this phase originating from precursor pyruvate (Zhou et al. 2010). Thus, the carbon atoms in n-alkyl lipids can originate from both assimilatory carbohydrates as well as transported carbohydrates from storage organs (Schmidt et al. 2003). Nonetheless, there are fractionations during lipid biosynthesis related to the numerous biochemical branch points which control the relationship between the δ13C of lipids, intermediates, and precursor molecules (Hayes 2001; Monson and Hayes 1980).

Coupling multiple isotope systems within a single plant tissue or compound can greatly expand the utility of isotope analysis (e.g., Mallette et al. 2017; Voelker et al. 2016). In this respect, linking the δ13C and δ15N values of plant tissues has increased the knowledge of carbon and nitrogen dynamics (e.g., Choi et al. 2005; Hobbie et al. 2017; Nasholm et al. 1998), while coupling δ13C and δ18O values from tree-ring cellulose has allowed for greater fidelity in the interpretation of climate archives from tree-ring record (e.g., Leavitt 2010; Szejner et al. 2016). Recently, the combination of δ13C, δ18O, and δ2H values of leaf cellulose was used to trace the transition from juvenile to mature period in leaf expansion and to demonstrate that hydrogen and oxygen in cellulose are controlled by different biochemistry (Kimak et al. 2015). The carbon and hydrogen isotope ratios of n-alkyl leaf wax lipids can be measured on the same individual compound with little to no additional purification or preparation. As n-alkyl leaf wax lipids are refractory in many geologic settings and can be isolated from soils, sediments, and rocks with straightforward procedures, the utility of these compounds is further amplified, as deeper time applications are possible. Strong relationships between a plant’s carbon metabolism, plant functional type, and the δ2H of n-alkyl lipids have been recognized (Chikaraishi and Naraoka 2007; Chikaraishi et al. 2004a; Cormier et al. 2018; Gamarra et al. 2016; Pedentchouk et al. 2008; Zhou et al. 2016), yet the majority of ancient and modern controlled studies on leaf wax lipids focus on either δ2H or δ13C values of these compounds exclusively. These concepts are just beginning to be applied to reconstruct paleoenvironments from leaf wax lipids isotope ratios (Magill et al. 2013; Tipple 2013; Tipple and Pagani 2010).

The mechanistic relationship between δ2H and δ13C values in n-alkyl lipids and how these values change during foliar development remains nascent. In broad-leaf species, variations in the δ2H of n-alkanes have been shown to relate to periods of leaf development (Freimuth et al. 2017; Kahmen et al. 2011; Oakes and Hren 2016; Sachse et al. 2015; Tipple et al. 2013). Pedentchouk et al. (2008) measured both δ2H and δ13C values of n-alkanes from broad-leafed species throughout a season and showed variations in both δ2H and δ13C of n-alkanes during the study period; however, these isotopic changes were not monitored in relation to foliar development. Recently, several studies have compared the δ2H of n-alkanes to δ13C of bulk foliar material through the growing season and found that both the n-alkanes and structural carbon were synthesized from stored and recent assimilate in the initial and later growth periods, respectively (Freimuth et al. 2017; Newberry et al. 2015). However, the relationships between δ2H and δ13C of n-alkanes in context to leaf development have not yet been explored.

Here, we investigate the multiple carbon and water sources that can influence the isotopic composition of n-alkanes in leaves. We report the δ2H and δ13C values of n-alkanes, the δ2H values of leaf, xylem, atmospheric, and stream waters, and the δ13C values of bulk foliar material, along with n-alkane molecular distributions from three common riparian tree species: Betula occidentalis, Populus angustifolia, and Acer negundo. Materials from these species were collected in a natural montane ecosystem, at 15 intervals throughout the growing season. Materials were specifically collected from leaves that developed during the initial leaf flush. These species and ecosystem were selected, as the δ2H value of stream water was known to remain relatively constant throughout the year and allowed for the effects of humidity and the δ2H of atmospheric water vapor on the δ2H of n-alkanes to be isolated. Concurrent collections of the δ2H from n-alkanes, xylem, leaf, and stream water allowed us to monitor the relationship between xylem and leaf water on the biosynthetic fractionation of n-alkanes. Also, the paired collections of δ13C values of n-alkanes and bulk leaf tissue provided information into the physiological and synthetic processes that connect these components of the leaf. Finally, the paired measurements of δ2H and δ13C of n-alkanes allowed us to evaluate how carbon metabolism and water environment are recorded within the same plant molecule, while jointly studying the effects of leaf ontogeny on these isotope systems. With this study, we sought to understand the temporal dynamics of coupled measurements of the δ2H and δ13C of n-alkanes and to determine the relationship between these isotope signatures and the heterotrophic and autotrophic processes that occur during foliar development and leaf expansion.

Materials and methods

Sample location and material collection

Materials from three individuals of Betula occidentalis Hook. (Western river birch), three individuals of Populus angustifolia James (narrowleaf cottonwood), and three individuals of Acer negundo L. (boxelder) were collected from a riparian environment near the Mill B South Fork of Big Cottonwood Creek in Big Cottonwood Canyon, Utah. Materials were collected from the same individuals on 15 dates throughout the 2010 Northern Hemisphere-growing season. Leaf samples were collected from the four cardinal directions from each individual and combined to reduce sun/shade variations. From each individual, approximately 4–8 buds or leaves were collected, combined, and stored in paper coin envelopes for leaf lipid and bulk carbon isotope analysis. The date on which the first full immature leaves were observed was described as “leaf flush” and occurred on DOY 141 for B. occidentalis and A. negundo, and DOY 146 for P. angustifolia. The date on which the first mature leaves were observed was noted as the mature phenophase and occurred on DOY 167 for B. occidentalis and A. negundo, and DOY 161 for P. angustifolia. The period including and between the “leaf flush” and the mature phenophase was termed the leaf expansion phenophase. The interval prior to “leaf flush” was described as the bud phenophase. Particular care was taken to only sample mature leaves from the initial leaf flush. These samples were dried for 48 h in a 50 °C oven immediately following collection.

Additional leaf samples were collected from the four cardinal directions of a single representative individual of the species above for water extraction. These samples were collected from the same representative of each of the three species throughout the season. Here, 4 leaves were collected and amalgamated in 4-ml ashed glass vials for leaf water analysis. Stem samples were collected from the same individual that was sampled for leaf water analysis. Woody stems from the first branch point of the tree’s trunk were collected and stored in 4-ml ashed glass vials. All leaf and stem water sample vials were capped, sealed with Parafilm®, and stored in a freezer until the time of processing.

Atmospheric water vapor was sampled using a cryo-trap following Helliker et al. (2002). Vapor samples were sampled on all collection days with the lone exception of the initial collection day. Briefly, atmospheric vapor was pumped through a glass cold trap submerged in a dry ice/ethanol slurry. After 1 h, the frozen atmospheric vapor was thawed and transferred to an ashed 4-ml vial, capped, and sealed with Parafilm®. The vial was stored in a freezer until the time of processing.

Water from Big Cottonwood Creek was sampled on each collection day. Water samples were collected in 4-ml ashed glass vials after rinsing the vial three times with water. Following collection, the vial was capped, sealed with Parafilm®, and stored in the freezer until the time of processing.

Plant water extraction and stable hydrogen isotope analysis of waters

Plant waters were extracted using a cryogenic vacuum water extraction method following West et al. (2006). After a complete extraction, the frozen plant water was thawed and transferred to an ashed 4-ml vial, capped, and sealed with Parafilm®. The vial was stored in a freezer until further processing. Prior to analysis, activated charcoal was added to plant waters to remove water-soluble organic compounds. After 48 h, the charcoal was removed and the water was transferred to ashed 2-ml autosampler vial. The 2-ml vial was stored in an 8 °C cooler until the time of analysis.

The stable hydrogen isotope ratios of plant waters, atmospheric water vapor, and stream water were analyzed on a Picarro model L1102-i water analyzer. Each water sample was analyzed four times (four consecutive replicate injections) with only the last two analyses being used. Unknowns were analyzed alongside a set of three in-house reference materials that were analyzed after every sixth unknown in an analytical sequence. The hydrogen isotope ratios of plant were normalized to the Vienna Standard Mean Ocean Water (VSMOW) scale using a two-point linear calibration of water primary reference materials, which had previously been standardized to the VSMOW scale [SIRFER Zero (0.1‰) and SIRFER DI (− 123.0‰)]. Precision for hydrogen isotope measurement of water was ± 2‰ (1σ, n = 40), as determined from secondary reference material [Evian, EV (− 72.5‰)].

All stable isotope ratios are reported in δ-notation:
$$\delta = \left[ {\left( {\frac{{R_{\text{sample}} }}{{R_{\text{std}} }}} \right) - 1} \right]$$
(1)
where R represents the 2H/1H or 13C/12C abundance ratio, and Rsample and Rstd are the ratios in the sample and standard, respectively. δ2H values are expressed relative to the standard VSMOW, while δ13C values are expressed relative to the standard Vienna Pee Dee Belemnite (VPDB).

Leaf preparation and stable carbon isotope analysis of bulk leaf material

Leaves were pulverized and homogenized in a solvent-washed ceramic mortar and pestle aided with liquid nitrogen. Ground leaf materials were transfer to an ashed 4-ml vial and stored until the time of processing. The stable carbon isotopes of bulk leaf material were analyzed using an EA 1110 CHN elemental analyzer (Carlo Erba, Milan, Italy) with a CONFLO III (Thermo Finnigan, Bremen, Germany) coupled to a Deltaplus isotope ratio mass spectrometer (Finnigan MAT, Bremen, Germany). Carbon isotope ratios of bulk plant material were normalized to the VPDB scale using a two-point linear calibration of primary reference materials, which had previously been standardized to the VPDB scale [UU-CN-1, glutamic acid (49.63‰) and UU-CN-2, glutamic acid (– 4.56‰)]. Primary reference materials were analyzed after every fifth unknown in an analytical sequence. Precision for carbon isotope measurement was ± 0.12‰ (1σ, n = 32), as determined from a secondary reference material [SIRFER Spinach (– 27.40‰)].

Lipid extraction, identification, and quantification

Lipids were extracted from 200 to 900 mg of ground leaf material with hexane by ultra-sonication (30 min × 2). The resulting extracts were concentrated under a gentle stream of purified nitrogen (N2) gas using a FlexiVap Work Station (Glas-Col, Terre Haute, IN, USA) and were transferred to 4-ml glass vials. Extracts were then purified by column chromatography using 1 g deactivated silica gel (70–230 mesh) in an ashed Pasteur pipette, and eluted with 2 ml hexane to obtain the saturated hydrocarbons following Tipple and Pagani (2013).

Saturated hydrocarbons were analyzed using a TraceGC gas chromatograph (GC) with a TraceMS mass spectrometer (Thermo Finnigan, Milan, Italy). The GC was equipped with a split-splitless injector at 300 °C and a fused silica, DB-5 phase column (30 m × 0.25 mm I.D., 0.25 μm film thickness; J&W Scientific, Agilent Technologies, Santa Clara, CA, USA) with helium as the carrier at a flow of 1.5 ml/min. The GC oven temperature program was 60–320 °C at 15 °C/min with an isothermal for 30 min. High molecular n-alkanes were identified through comparison of elution times with known n-alkane standards.

Chain length distributions and concentrations of n-alkanes

Peak areas of high molecular weight n-alkanes were measured to quantify concentration and distribution. Concentrations were quantified using a 4-point calibration curve generated from reference materials of known concentration. The carbon preference index (CPI) was calculated following Marzi et al. (1993):
$${\text{CPI}}\; = \;\frac{{(A_{23} + A_{25} + A_{27} + A_{29} + A_{31} + A_{33} ) + (A_{25} + A_{27} + A_{29} + A_{31} + A_{33} + A_{35} )}}{{2(A_{24} + A_{26} + A_{28} + A_{30} + A_{32} + A_{34} )}}$$
(2)
where “A” represents the area of the individual n-alkane peaks. The average chain length (ACL) of n-alkanes was calculated:
$${\text{ACL}}\; = \;\frac{{(A_{23} (23)) + (A_{25} (25)) + (A_{31} (27)) + (A_{29} (29)) + (A_{31} (31)) + (A_{33} (33)) + (A_{35} (35))}}{{(A_{23} + A_{25} + A_{27} + A_{29} + A_{31} + A_{33} + A_{35} )}}$$
(3)

Compound-specific hydrogen and carbon isotope analysis

Compound-specific hydrogen isotope analyses were performed at the SIRFER Laboratory, University of Utah using a Trace GC Ultra gas chromatograph (Thermo Electron Corporation, Milan, Italy) equipped with a split-splitless injector at 300 °C and a fused silica, DB-5 phase column (30 m × 0.25 mm I.D., 0.25 μm film thickness; J&W Scientific, Agilent Technologies, Santa Clara, CA, USA). Helium was the carrier, with a flow rate of 1.2 ml/min. The GC oven temperature program was 102–350 °C at 6 °C/min with an isothermal for 8 min. Following GC separation, n-alkanes were pyrolyzed at 1400 °C in a High Temperature Conversion system (Thermo Scientific, Bremen, Germany) and subsequently analyzed for hydrogen isotope ratios using a Delta V Plus isotope ratio mass spectrometer (Thermo Scientific, Bremen, Germany). The H3+ factor was determined daily prior to standard calibration and sample analysis. The average H3+ factor during these analyses was 2.42 ± 0.25 (1σ, n = 43). Individual n-alkane hydrogen isotope ratios were normalized to the VSMOW scale using a two-point linear calibration of n-alkane primary reference materials, which had previously been standardized to the VSMOW scale [n-C24 (– 36‰) and n-C28 (– 250‰)]. Primary reference materials were analyzed after every fifth unknown in an analytical sequence. Precision for n-alkanes hydrogen isotope determinations was ± 3‰ (1σ, n = 165), as determined from a secondary n-alkane reference material [n-C36 (– 240‰)]. The apparent fractionation between n-alkane and leaf water (εH) is defined as:
$$\varepsilon_{\text{H}} = \left[ {\left( {\frac{{R_{{n - {\text{alkane}}}} }}{{R_{\text{LW}} }}} \right) - 1} \right]$$
(4)
where R represents the 2H/1H abundance ratio, and Rn-alkane and RLW are the ratios in the n-alkane and leaf water, respectively.
Compound-specific carbon isotope analyses were also performed in the SIRFER Laboratory at the University of Utah. A 6890A GC (Hewlett-Packard Company, Palo Alto, CA, USA) employing a split–splitless injector held at a constant 310 °C with a fused silica, DB-5 phase column (30 m × 0.25 mm I.D., 0.25 μm film thickness; J&W Scientific, Agilent Technologies, Santa Clara, CA, USA) was used. Helium was the carrier, with a flow rate of 1.2 ml/min. The GC oven temperature program was 80–320 °C at 6 °C/min with an isothermal for 12 min. Compounds were subsequently combusted over nickel oxide, copper oxide, and platinum at 1000 °C using a GC Combustion III system (Thermo Quest Finnigan, Bremen, Germany) and analyzed for carbon isotope ratios using a DeltaplusXL isotope ratio mass spectrometer (Thermo Quest Finnigan, Bremen, Germany). Individual n-alkane carbon isotope ratios were normalized to the VPDB scale using a two-point linear calibration of n-alkane primary reference materials, which had previously been standardized to the VPDB scale [n-C18 (– 33.3‰) and n-C28 (– 29.0‰)]. Primary reference materials were analyzed after every fifth unknown in an analytical sequence. Arndt Schimmelmann’s “Mix A4” [n-C16 (– 30.7‰), n-C17 (– 31.2‰), n-C18 (– 31.1‰), n-C19 (–33.2‰), n-C20 (– 32.4‰), n-C21 (– 29.1‰), n-C22 (– 32.9‰), n-C23 (– 31.8‰), n-C24 (– 33.3‰), n-C25 (–28.5‰), n-C26 (– 33.0‰), n-C27 (– 29.6‰), n-C28 (–32.2‰), n-C29 (– 30.1‰), and n-C30 (– 29.9‰)] was analyzed twice in each analytical sequence (n =32) and had a measured accuracy of 0.1‰. Precision for n-alkanes carbon isotope determinations was ± 0.3‰ (1σ, n = 157), as determined from a co-injected secondary reference material [5α-androstane (– 30.1‰)]. The apparent fractionation between n-alkane and bulk leaf (εC) is defined as:
$$\varepsilon_{\text{C}} = \left[ {\left( {\frac{{R_{{n - {\text{alkane}}}} }}{{R_{\text{bulk}} }}} \right) - 1} \right]$$
(5)
where R represents the 13C/12C abundance ratio, and Rn-alkane and Rbulk are the ratios in the n-alkane and bulk leaf, respectively.

Statistical analysis

Statistical analysis was completed using JMP Pro 13® (SAS Institute Inc., Cary, NC, USA). The isotope data from a given sampling day, species, and leaf developmental stage were compared using Student t tests to identify differences (α < 0.05).

Results

Abundance and distribution of n-alkanes

The concentrations of individual n-alkane homologs and total n-alkanes relative to dry weight were determined (Table 1). Concentration data for each individual and all measured n-alkane homologs are reported in Table S1. In addition, the average chain length (ACL) values were calculated (Table 1). Here, we found that each species produced a unique distribution of n-alkanes with variations in distribution and concentration between the species and leaf developmental stage. Betula occidentalis produced n-C23 to n-C33 with n-C27, n-C29, and n-C31 being the most abundant homologs. The three B. occidentalis individuals had average concentrations of 243 ± 130, 89 ± 35, and 117 ± 44 µg g−1 over the sampling period for n-C27, n-C29, and n-C31, respectively (Table 1). The total concentration of n-alkanes on all B. occidentalis individuals ranged from 44 to 1899 µg g−1 over the sampling period with an average of 486 ± 510 µg g−1. In general, the total n-alkane concentration in B. occidentalis decreased 10-fold from the bud phenophase through leaf expansion (DOY 112 to 167) and then increased slightly during leaf maturation (DOY 179 to 239) and then decreased subsequently (Fig. 1). However, the relative distribution of n-alkanes varied throughout the leaf developmental stages with the n-C29/n-C31 ratio varying from 2.04 to 8.94 in the bud to leaf flush (DOY 112 to 141) and then dropping precipitously to an average of 0.36 in the mature leaf (Table S1).
Table 1

Average concentrations of individual and total n-alkanes and molecular distrtibutions of n-alkanes from three riparian tree species

Ordinal date

Phenophase

[n-C27] (mg g−1)

 

SD

[n-C29] (mg g−1)

 

SD

[n-alkane] (mg g−1)

 

SD

CPI

 

SD

ACL

 

SD

Betula occidentalis

 112

Bud

0.8807

±

0.7449

0.2780

±

0.1742

1.341

±

1.067

35.6

±

9.7

27.68

±

0.46

 123

Bud

1.2350

±

0.5486

0.3511

±

0.0287

1.899

±

0.621

41.9

±

11.3

27.64

±

0.41

 132

Bud

0.4186

±

0.1430

0.1076

±

0.0140

0.663

±

0.230

27.0

±

1.7

27.23

±

0.48

 137

Bud

0.2141

±

0.1351

0.0415

±

0.0211

0.327

±

0.225

46.7

±

19.7

26.87

±

0.48

 141

Leaf expansion

0.1223

±

0.0378

0.0210

±

0.0013

0.181

±

0.055

29.6

±

0.5

26.74

±

0.39

 146

Leaf expansion

0.0849

±

0.0342

0.0102

±

0.0026

0.139

±

0.062

35.0

±

4.8

26.28

±

0.46

 153

Leaf expansion

0.0995

±

0.0507

0.0127

±

0.0048

0.166

±

0.091

23.8

±

5.9

26.35

±

0.51

 161

Leaf expansion

0.0843

±

0.0249

0.0748

±

0.0932

0.213

±

0.120

45.4

±

27.0

27.07

±

1.24

 167

Leaf expansion

0.0192

±

0.0160

0.0063

±

0.0057

0.044

±

0.031

20.4

±

3.2

26.90

±

1.49

 179

Mature

0.0526

±

0.0324

0.0246

±

0.0028

0.162

±

0.079

17.7

±

2.3

28.35

±

0.98

 195

Mature

0.0683

±

0.0275

0.0538

±

0.0180

0.285

±

0.035

26.7

±

1.5

29.01

±

0.98

 216

Mature

0.1334

±

0.0592

0.1412

±

0.0829

0.703

±

0.246

22.5

±

2.0

29.54

±

0.71

 239

Mature

0.1105

±

0.0402

0.1092

±

0.0264

0.577

±

0.072

18.9

±

0.8

29.73

±

0.40

 250

Mature

0.0555

±

0.0146

0.0509

±

0.0248

0.276

±

0.069

18.7

±

0.7

29.75

±

0.22

 274

Mature

0.0622

±

0.0246

0.0570

±

0.0189

0.313

±

0.118

18.0

±

3.4

29.73

±

0.29

Populus angustifolia

 112

Bud

0.3485

±

0.0630

0.1069

±

0.0085

0.515

±

0.062

24.5

±

9.5

27.36

±

0.15

 123

Bud

0.4687

±

0.3604

0.1339

±

0.1160

0.718

±

0.621

29.0

±

11.0

27.30

±

0.13

 132

Bud

0.4851

±

0.0283

0.1586

±

0.0068

0.741

±

0.034

23.1

±

8.4

27.37

±

0.12

 137

Bud

0.2815

±

0.0628

0.1003

±

0.0276

0.431

±

0.115

26.9

±

14.6

27.39

±

0.09

 141

Bud

0.3292

±

0.1118

0.1392

±

0.0395

0.519

±

0.164

25.8

±

7.1

27.50

±

0.01

 146

Leaf expansion

0.3156

±

0.1007

0.3379

±

0.3288

0.699

±

0.446

32.1

±

9.8

27.81

±

0.44

 153

Leaf expansion

0.3971

±

0.1127

0.9449

±

0.3303

1.447

±

0.454

21.5

±

1.8

28.42

±

0.15

 161

Leaf expansion

0.1758

±

0.1116

1.2525

±

0.4393

1.492

±

0.590

34.5

±

8.1

28.81

±

0.09

 167

Mature

0.0985

±

0.0227

1.1023

±

0.3261

1.251

±

0.277

46.8

±

48.8

28.86

±

0.06

 179

Mature

0.0955

±

0.0316

0.5549

±

0.1473

0.705

±

0.184

23.4

±

12.2

28.78

±

0.04

 195

Mature

0.1110

±

0.0024

0.6711

±

0.1524

0.840

±

0.145

31.7

±

7.1

28.76

±

0.04

 216

Mature

0.1228

±

0.0243

0.7676

±

0.0360

0.965

±

0.054

25.6

±

3.4

28.95

±

0.32

 239

Mature

±

±

±

±

±

 250

Mature

0.0513

±

0.0241

0.4917

±

0.1416

0.579

±

0.187

30.9

±

21.9

28.82

±

0.06

 274

Mature

0.0669

±

0.0059

0.5335

±

0.0399

0.644

±

0.032

23.9

±

6.7

28.77

±

0.05

Acer negundo

 112

Bud

0.2798

±

0.1458

1.9187

±

0.2296

3.876

±

0.376

4.8

±

0.6

30.67

±

0.41

 123

Bud

0.1562

±

0.1436

0.2700

±

0.3185

0.873

±

0.302

3.7

±

3.6

29.62

±

0.60

 132

Bud

0.0810

±

0.0558

0.3696

±

0.2916

0.878

±

0.436

5.4

±

4.7

29.54

±

0.51

 137

Bud

±

±

±

±

±

 141

Leaf expansion

0.0957

±

0.0234

0.3959

±

0.0840

0.824

±

0.276

7.0

±

2.1

29.85

±

0.59

 146

Leaf expansion

0.0867

±

0.0074

0.3799

±

0.0769

0.733

±

0.156

5.7

±

0.8

30.00

±

0.21

 153

Leaf expansion

0.0683

±

0.0118

0.2683

±

0.0794

0.559

±

0.154

5.5

±

1.7

29.91

±

0.21

 161

Leaf expansion

0.1632

±

0.0462

0.5590

±

0.0916

1.177

±

0.249

5.1

±

1.1

30.23

±

0.08

 167

Leaf expansion

0.1674

±

0.0358

0.5656

±

0.1444

1.177

±

0.243

5.9

±

1.4

30.24

±

0.05

 179

Mature

0.0743

±

0.0347

0.3806

±

0.1476

0.645

±

0.268

9.7

±

0.9

30.62

±

0.02

 195

Mature

0.1335

±

0.1229

0.6722

±

0.5655

1.116

±

0.962

12.0

±

2.6

30.60

±

0.08

 216

Mature

0.0719

±

0.0121

0.3951

±

0.0986

0.620

±

0.124

10.8

±

1.6

30.61

±

0.11

 239

Mature

0.1968

±

0.1974

0.8849

±

0.7935

1.440

±

1.399

11.9

±

1.7

30.50

±

0.18

 250

Mature

0.0492

±

0.0087

0.3534

±

0.1428

0.516

±

0.189

13.2

±

2.2

30.71

±

0.17

 274

Mature

0.0574

±

0.0117

0.2891

±

0.0738

0.467

±

0.076

10.0

±

2.9

30.50

±

0.32

Fig. 1

Concentration of n-C27, n-C29, and n-C31 alkanes on buds and leaves of a Betula occidentalis, b Populus angustifolia, and c Acer negundo. Concentration data from n-C27, n-C29, and n-C31 are shown in closed, open, and crossed squares. Error bars represent the standard deviations around the mean of three individuals from each species. Average chain length of n-alkane from bud and leaf waxes, d B. occidentalis, e P. angustifolia, and f A. negundo. Error bars represent the standard deviations around the mean of the three individuals from each species used in this study. Colored areas correspond to the bud (white), leaf expansion (dark gray), and mature leaf (light gray) phenophases

Populus angustifolia produced n-C23 to n-C31 with n-C27 and n-C29 being the most abundant homologs. In the three P. angustifolia individuals sampled, n-C29 (521 ± 153 µg g−1) was the most abundant homolog on average, approximately two times more abundant than the next most abundant homolog, n-C27 (239 ± 76 µg g−1) (Table 1). The average total concentration of n-alkanes on P. angustifolia individuals was 825 ± 342 µg g−1. The total n-alkane concentrations in P. angustifolia were distinct from the other species and increased over from 43 to 1492 µg g−1 during leaf expansion (DOY 147 to 161) and then decreased during the mature phenophase (Fig. 1). However, the relative distribution of n-alkanes varied in P. angustifolia throughout the leaf developmental stages with the n-C27/n-C29 ratio varying from an average value of 2.83 in the bud phenophase to leaf flush (DOY 112 to 146) to an average 0.14 in the mature leaf (Table S1).

The composition of n-alkanes on Acer negundo was n-C23 to n-C33, with n-C31 as the most abundant and with n-C27 and n-C29 being roughly five-times less concentrated than n-C31. The three A. negundo individuals sampled had an average n-C31 concentration of 550 ± 224 µg g−1. A. negundo had an average total concentration of n-alkanes 1064 ± 224 µg g−1 with a range of 467 to 3876 µg g−1. The total n-alkane concentration in A. negundo decreased significantly during the bud to leaf flush (DOY 112 to 141) and then remained relatively stable throughout the leaf expansion and mature leaf phenophases (Fig. 1). While the total n-alkane concentration remained stable, the relative distribution subtly varied across the season. The C27/n-C29 ratio generally decreased from the initial leaf expansion phenophase to the mature leaf phenophase, while the n-C29/n-C31 remained relatively stable during this period (Table 1).

We found that all species showed systematic changes in the average chain length (ACL) during the season and leaf developmental phenophases. Here, all species had ACL values that remained stable or decreased from bud to leaf flush (Fig. 1). The ACL of all species increased during the leaf expansion phenophase. ACL stabilized once the leaves of P. angustifolia and A. negundo matured, while the ACL of B. occidentalis continued to increase until DOY 239, after which it stabilized. B. occidentalis individuals had an average ACL of 27.9 ± 1.3 with a range from 26.3 to 29.8. P. angustifolia individuals had an average ACL of 28.2 ± 0.7 with a range from 27.3 to 28.9. Finally, A. negundo individuals had an average ACL of 30.3 ± 0.4 with a range from 29.5 to 30.7.

Hydrogen isotopes of plant and environmental waters

The hydrogen isotope values of stream water and atmospheric water vapor are presented in Table 2. The mean hydrogen isotope value of stream water (δ2HMW) was − 130 ± 1‰ (1σ, n = 15; Table 2). The δ2HMW of Big Cottonwood Creek waters did not vary significantly during the studied interval as the major source of water derived from upper elevation snow melt (Roden and Ehleringer 2000). The mean hydrogen isotope value of atmospheric water vapor (δ2HV) was − 172 ± 24‰ (1σ, n = 14) and ranged from − 233 to − 132‰. The δ2HV varied over 100‰ due to changes in moisture source regions. Vapor was most 2H-depleted early in the growing season, reflecting cooler air masses originating from the Northern Pacific.
Table 2

Hydrogen isotope values of Big Cottonwood Creek water, atmospheric water vapors as well as xylem and leaf water from three riparian tree species

Ordinal date

δ2H stream (‰, VSMOW)

δ2H vapor (‰, VSMOW)

Betula occidentalis

Populus angustifolia

Acer negundo

Phenophase

δ2H stem (‰, VSMOW)

 

SD

δ2H leaf (‰, VSMOW)

 

SD

Phenophase

δ2H stem (‰, VSMOW)

 

SD

δ2H leaf (‰, VSMOW)

 

SD

Phenophase

δ2H stem (‰, VSMOW)

 

SD

δ2H leaf (‰, VSMOW)

 

SD

112

− 132.2

Bud

− 118.2

±

0.0

− 107.4

±

Bud

− 123.1

±

0.3

±

Bud

− 116.8

±

0.0

− 157.3

±

0.9

123

− 131.2

− 172.9

Bud

− 100.3

±

0.3

− 100.6

±

0.5

Bud

− 117.8

±

0.2

− 93.2

±

0.0

Bud

− 86.9

±

0.4

− 80.6

±

2.2

132

− 131.5

− 233.4

Bud

− 113.1

±

0.0

− 109.4

±

0.2

Bud

− 110.2

±

1.1

− 99.6

±

1.2

Bud

− 89.3

±

0.2

− 87.0

±

1.1

137

− 131.2

− 185.5

Bud

− 119.0

±

0.0

− 116.6

±

Bud

− 132.0

±

0.2

− 100.4

±

Bud

− 109.5

±

0.1

− 73.2

±

0.5

141

− 130.5

− 157.5

Leaf expansion

− 107.2

±

− 84.7

±

2.1

Bud

− 120.7

±

0.1

− 85.2

±

0.1

Leaf expansion

− 101.9

±

0.4

− 82.1

±

0.4

146

− 130.1

− 159.6

Leaf expansion

− 114.5

±

0.7

− 99.6

±

0.3

Leaf expansion

− 120.4

±

0.7

− 93.1

±

0.5

Leaf expansion

− 108.3

±

− 71.1

±

0.0

153

− 129.4

− 187.6

Leaf expansion

− 120.7

±

− 122.4

±

1.0

Leaf expansion

− 127.1

±

0.2

− 115.7

±

0.5

Leaf expansion

− 114.7

±

0.8

− 105.6

±

0.5

161

− 127.9

− 168.9

Leaf expansion

− 130.8

±

0.9

− 99.7

±

Leaf expansion

− 135.8

±

0.1

− 96.1

±

0.4

Leaf expansion

− 130.4

±

0.2

− 95.3

±

0.3

167

− 129.3

− 169.0

Leaf expansion

− 133.1

±

0.2

− 87.2

±

0.2

Mature

− 133.4

±

0.6

− 114.7

±

1.1

Leaf expansion

− 132.4

±

− 99.1

±

0.8

179

− 130.2

− 186.5

Mature

− 139.4

±

0.3

− 98.4

±

Mature

− 133.5

±

0.1

− 111.8

±

0.6

Mature

− 133.7

±

0.1

− 132.1

±

0.0

195

− 129.9

− 167.8

Mature

− 135.8

±

1.4

− 85.4

±

Mature

− 133.9

±

0.1

− 75.9

±

Mature

− 130.6

±

0.2

− 74.7

±

0.5

216

− 129.6

− 139.0

Mature

− 136.3

±

− 81.9

±

0.5

Mature

− 131.2

±

− 68.4

±

1.1

Mature

− 122.5

±

0.5

− 66.6

±

0.8

239

− 129.0

− 132.3

Mature

− 139.9

±

0.2

− 59.0

±

1.2

Mature

− 133.3

±

0.9

− 60.6

±

0.5

Mature

− 123.5

±

0.3

− 41.6

±

0.2

250

− 129.8

− 168.9

Mature

− 133.3

±

0.3

− 65.2

±

0.3

Mature

− 136.1

±

0.7

− 69.1

±

0.9

Mature

− 120.1

±

0.4

− 38.3

±

0.2

274

− 127.5

− 176.1

Mature

− 140.6

±

0.4

− 56.2

±

Mature

− 145.3

±

0.8

− 61.1

±

0.4

Mature

− 119.2

±

0.1

− 39.5

±

0.0

The hydrogen isotope values of stem and leaf waters are provided in Table 2 and shown in Fig. 2. Here, we found that hydrogen isotope values of stem waters (δ2HXW) in all species were more 2H-enriched values in the initial growing season and became relatively more 2H-depleted prior to leaf maturity. The δ2HXW from all species ranged from − 87 to − 145‰ (Table 2). Mean δ2HXW of B. occidentalis, P. angustifolia, and A. negundo was − 125 ± 13, − 129 ± 9, and − 116 ± 15‰ (1σ, n = 15 for each species), respectively. Mean δ2HXW of B. occidentalis and P. angustifolia was not different, while δ2HXW from A. negundo was significantly different from the other two species (t test, p = 0.0191 and 0.0017, respectively). Following the bud and leaf expansion phenophases, the δ2HXW for all species stabilized and remained relatively constant for the remainder of the growing season (Fig. 2). Mean δ2HXW of B. occidentalis, P. angustifolia, and A. negundo during the mature leaf phenophase was − 138 ± 3‰ (1σ, n = 6), − 135 ± 5‰ (1σ, n = 7), and − 125 ± 6‰ (1σ, n = 6), respectively. Additionally, the δ2HXW of B. occidentalis and P. angustifolia of each phenophase was not significantly different, while the δ2HXW of A. negundo was significantly different from P. angustifolia during the bud phenophase (t test, p = 0.0182) and both B. occidentalis and P. angustifolia during the mature leaf phenophase (t-test, p = 0.0002 and 0.0010, respectively). In addition, the pooled mean δ2HXW of all species during the mature phenophase was not significantly different than δ2HMW; however, the mean δ2HXW of each individual species during the mature phenophase was significantly different than δ2HMW (t test, all p < 0.006).
Fig. 2

Hydrogen isotope values of stem water, leaf water, and n-alkanes from a Betula occidentalis, b Populus angustifolia, and c Acer negundo during the studied interval. Hydrogen isotope values of stem and leaf water are shown in closed and open circles, while data from n-C27, n-C29, and n-C31 are shown in closed, open, and crossed squares, respectively. Error bars represent the standard deviations around the mean of three P. angustifolia specimens. Colored areas correspond to the bud (white), leaf expansion (dark gray), and mature leaf (light gray) phenophases

The hydrogen isotope values of leaf waters (δ2HLW) varied considerably during the growing season, ranging from − 38 to − 157‰. Mean δ2HLW of B. occidentalis, P. angustifolia, and A. negundo was − 92 ± 20‰ (1σ, n = 15), − 89 ± 19‰ (1σ, n = 14), and − 83 ± 33‰ (1σ, n = 15), respectively. The overall mean δ2HLW was not different among species, nor was the mean δ2HLW for each leaf developmental phenophase. Generally, δ2HLW and δ2HXW were relatively similar during the bud and leaf expansion phenophases, with δ2HLW being slightly less negative, while as the season progressed, δ2HLW became increasingly less negative compared to δ2HXW (Fig. 2). The difference (Δ) between δ2HLW and δ2HXW was 9 ± 21‰ (1σ, n = 10), 20 ± 13‰ (1σ, n = 12), and 56 ± 23‰ (1σ, n = 22) for the bud, leaf expansion, and mature phenophases, respectively. The δ2HLW was positively correlated with δ2HV (R2 = 0.72, p < 0.0001) from leaf maturity until DOY 239.

Hydrogen isotopes of n-alkanes and fractionations between n-alkanes and leaf water

The hydrogen isotope values of prominent n-alkane homologs are reported in Table 3 and Fig. 2. Here, the δ2H of n-C27 and n-C29 was measured in B. occidentalis, while it was only possible to measure n-C31 in A. negundo. While hydrogen isotope analysis was conducted on all homologs of sufficient abundance, we report and discuss the hydrogen isotope values of only the most abundant homolog for each species (δ2Hn-alkane). Nonetheless, the δ2H of n-C27 and n-C29 from B. occidentalis was highly correlated (R2 = 0.93, p < 0.0001). The δ2H of n-C27 and n-C29 from P. angustifolia has been previously published (Tipple et al. 2013) and will not be extensively described here.
Table 3

Hydrogen isotope values of n-alkanes from the three species

Ordinal Date

Betula occidentalis

Populus angustifolia

Acer negundo

Phenophase

δ2H n-C27 (‰, VPDB)

 

SD

δ2H n-C29 (‰, VPDB)

 

SD

Phenophase

δ2H n-C27 (‰, VPDB)

 

SD

δ2H n-C29 (‰, VPDB)

 

SD

Phenophase

δ2H n-C29 (‰, VPDB)

 

SD

δ2H n-C31 (‰, VPDB)

 

SD

112

Bud

− 269.5

±

n/a

− 274.9

±

n/a

Bud

− 269.0

±

6.5

− 267.5

±

10.6

Bud

− 216.2

±

n/a

− 249.6

±

n/a

123

Bud

− 265.0

±

n/a

− 267.9

±

n/a

Bud

− 267.7

±

1.3

− 269.3

±

2.9

Bud

±

n/a

− 229.5

±

16.2

132

Bud

− 235.8

±

n/a

− 247.1

±

n/a

Bud

− 262.8

±

5.4

− 260.9

±

2.6

Bud

− 193.6

±

n/a

− 217.1

±

7.1

137

Bud

− 238.3

±

n/a

− 253.7

±

n/a

Bud

− 257.9

±

0.3

− 246.2

±

1.4

Bud

− 202.6

±

n/a

− 214.6

±

7.4

141

Leaf expansion

− 238.0

±

n/a

− 243.7

±

n/a

Bud

− 238.5

±

7.1

− 226.9

±

10.0

Leaf expansion

− 194.5

±

n/a

− 216.4

±

5.5

146

Leaf expansion

− 217.3

±

n/a

− 229.9

±

n/a

Leaf expansion

− 223.7

±

2.5

− 210.8

±

5.2

Leaf expansion

±

n/a

− 216.8

±

3.2

153

Leaf expansion

− 216.2

±

n/a

− 227.8

±

n/a

Leaf expansion

− 213.5

±

4.7

− 207.8

±

2.2

Leaf expansion

− 192.6

±

n/a

− 213.8

±

0.7

161

Leaf expansion

− 209.6

±

n/a

− 213.6

±

n/a

Leaf expansion

− 217.8

±

11.1

− 221.5

±

3.8

Leaf expansion

− 198.6

±

n/a

− 216.8

±

6.1

167

Leaf expansion

− 203.6

±

n/a

− 211.5

±

n/a

Mature

− 217.1

±

3.5

− 224.9

±

3.3

Leaf expansion

±

n/a

− 212.6

±

4.3

179

Mature

− 190.7

±

n/a

− 187.0

±

n/a

Mature

− 209.5

±

1.8

− 215.3

±

2.6

Mature

±

n/a

− 211.7

±

3.9

195

Mature

− 197.3

±

n/a

− 200.0

±

n/a

Mature

− 213.9

±

5.2

− 218.8

±

2.1

Mature

±

n/a

− 214.9

±

4.1

216

Mature

− 196.5

±

n/a

− 189.2

±

n/a

Mature

− 209.4

±

4.5

− 217.6

±

3.6

Mature

±

n/a

− 212.7

±

5.3

239

Mature

− 208.2

±

n/a

− 195.9

±

n/a

Mature

− 211.8

±

8.1

− 222.8

±

5.4

Mature

− 206.2

±

n/a

− 217.9

±

n/a

250

Mature

− 199.6

±

n/a

− 199.3

±

n/a

Mature

− 209.3

±

9.9

− 219.4

±

1.8

Mature

− 214.9

±

n/a

− 214.5

±

1.1

274

Mature

− 205.8

±

n/a

− 197.5

±

n/a

Mature

− 215.1

±

8.0

− 217.2

±

4.0

Mature

− 202.3

±

n/a

− 215.0

±

6.7

During specific phenophases, distinct patterns in the δ2Hn-alkane of B. occidentalis and A. negundo were observed. In both species, the initial δ2Hn-alkane values during the bud phenophase were the most negative and became less negative prior to leaf flush. A similar pattern was observed in P. angustifolia. During the transition from bud to leaf flush, the δ2Hn-alkane of B. occidentalis, P. angustifolia, and A. negundo became more enriched by 20, 57, and 35‰, respectively. During leaf expansion, δ2Hn-alkane of B. occidentalis continued to became more 2H-enriched by 42‰, while the δ2Hn-alkane of A. negundo stabilized, becoming only 2‰ more 2H-enriched. In comparison, P. angustifolia became more 2H-depleted by 11‰. From bud through leaf expansion, the total enrichment in 2H for B. occidentalis (DOY 112 to 167), P. angustifolia (DOY 112 to 161), and A. negundo (DOY 112 to 167) was 63, 46, and 37‰, respectively. The enrichment pattern in δ2Hn-alkane corresponding to the same interval in that δ2HXW became more 2H-depleted. After the leaf expansion phenophase, the δ2Hn-alkane values became very stable, where the mean δ2Hn-alkane value remained at − 195 ± 5‰ (1σ, n = 6), − 219 ± 3‰ (1σ, n = 7), and − 214 ± 2‰ (1σ, n = 6) for B. occidentalis, P. angustifolia, and A. negundo, respectively.

The apparent hydrogen isotope fractionation (εH) between δ2Hn-alkane and δ2HLW, shown in Fig. 3, varied systematically across the growing season for all species. In general, we found that the initial εH values became relatively less negative from the bud phenophase through the leaf expansion phenophase. After leaf expansion, the εH values became more negative. The period from leaf flush to full leaf expansion is known to be the interval of wax synthesis in broad-leafed angiosperms (Freimuth et al. 2017; Kahmen et al. 2011; Oakes and Hren 2016; Tipple et al. 2013). During this interval, we found that the pooled mean εH value for all species was − 132 ± 18‰ (1σ, n = 13) and the individual mean εH values were − 131 ± 22‰ (1σ, n = 5), − 124 ± 17‰ (1σ, n = 3), − 137 ± 15‰ (1σ, n = 5) for B. occidentalis, P. angustifolia, and A. negundo, respectively. Following the leaf expansion and wax synthesis, εH values for all species gradually became more negative due to the 2H-enrichment in the leaf water (Fig. 3).
Fig. 3

Apparent fractionation between the hydrogen isotope values of leaf water and normal alkanes from Betula occidentalis, Populus angustifolia, and Acer negundo during the studied period

Carbon isotopes of n-alkanes and bulk leaf materials

The carbon isotope values of bulk leaf material (δ13Cbulk) for all species are reported in Table 4 and Fig. 4. Within all species, a defined 13C-enrichment was observed in δ13Cbulk values during the bud and leaf expansion phenophases. Here, all species showed a slight 13C-depletion, followed by a 13C-enrichment prior to leaf flush (Fig. 4). The least negative δ13Cbulk values were observed at or immediately following leaf flush, with values of − 28.0, − 29.5, and − 27.7‰ for B. occidentalis, P. angustifolia, and A. negundo, respectively. δ13Cbulk subsequently became more negative during the leaf expansion phenophase by 2.1, 0.4, and 1.6‰ for B. occidentalis, P. angustifolia, and A. negundo, respectively. Following leaf expansion, the δ13Cbulk in all species remained stable through the growing season, where the mean δ13Cbulk was − 29.7 ± 0.3‰ (1σ, n = 6), − 31.3 ± 0.3‰ (1σ, n = 7), and − 30.1 ± 0.2‰ (1σ, n = 6) for B. occidentalis, P. angustifolia, and A. negundo, respectively.
Table 4

Carbon isotope values of bulk foliar material and mean n-alkanes from the three tree species

Ordinal date

Betula occidentalis

Populus angustifolia

Acer negundo

Phenophase

δ13C bulk leaf (‰, VPDB)

 

SD

δ13C n-C27 (‰, VPDB)

 

SD

δ13C n-C29 (‰, VPDB)

 

SD

Phenophase

δ13C bulk leaf (‰, VPDB)

 

SD

δ13C n-C27 (‰, VPDB)

 

SD

δ13C n-C29 (‰, VPDB)

 

SD

Phenophase

δ13C bulk leaf (‰, VPDB)

 

SD

δ13C n-C29 (‰, VPDB)

 

SD

δ13C n-C31 (‰, VPDB)

 

SD

112

Bud

− 28.7

±

1.0

− 31.9

±

2.0

− 31.6

±

1.6

Bud

− 31.1

±

0.9

− 35.6

±

0.9

− 34.8

±

0.8

Bud

− 27.2

±

1.3

− 33.9

±

1.1

− 33.2

±

0.7

123

Bud

− 29.2

±

1.3

− 32.7

±

2.6

− 32.6

±

2.3

Bud

− 30.8

±

1.2

− 35.6

±

1.3

− 34.5

±

1.4

Bud

− 28.4

±

1.2

− 33.3

±

1.3

− 32.7

±

1.7

132

Bud

− 28.8

±

0.8

− 33.5

±

2.4

− 33.5

±

2.4

Bud

− 31.3

±

0.5

− 36.2

±

0.6

− 35.4

±

0.5

Bud

− 28.3

±

0.8

− 33.5

±

0.6

− 32.7

±

0.5

137

Bud

− 28.0

±

0.9

− 33.1

±

1.5

− 33.1

±

1.3

Bud

− 30.9

±

1.4

− 36.0

±

1.4

− 35.5

±

1.4

Bud

− 27.7

±

0.7

− 33.3

±

1.2

− 32.6

±

1.6

141

Leaf expansion

− 28.0

±

0.8

− 33.7

±

2.0

− 33.6

±

1.9

Bud

− 30.1

±

0.3

− 36.5

±

0.1

− 36.2

±

0.2

Leaf expansion

− 28.2

±

0.7

− 33.5

±

0.7

− 32.6

±

0.5

146

Leaf expansion

− 27.9

±

0.6

− 34.1

±

0.9

− 34.6

±

1.1

Leaf expansion

− 29.5

±

1.1

− 36.5

±

0.8

− 36.5

±

0.5

Leaf expansion

− 28.3

±

1.0

− 33.8

±

1.3

− 33.0

±

1.3

153

Leaf expansion

− 28.2

±

0.6

− 32.9

±

1.8

− 32.7

±

1.8

Leaf expansion

− 29.9

±

0.7

− 36.6

±

0.5

− 36.2

±

0.6

Leaf expansion

− 28.3

±

0.7

− 34.1

±

0.7

− 33.4

±

0.6

161

Leaf expansion

− 29.4

±

0.7

− 33.6

±

1.4

− 34.4

±

1.1

Leaf expansion

− 30.3

±

0.1

− 36.0

±

1.0

− 34.1

±

1.2

Leaf expansion

− 29.6

±

1.0

− 35.1

±

1.7

− 35.1

±

1.5

167

Leaf expansion

− 30.1

±

0.9

− 35.0

±

1.5

− 34.6

±

1.0

Mature

− 31.1

±

0.4

− 36.3

±

0.1

− 34.5

±

0.5

Leaf expansion

− 29.8

±

0.6

− 36.2

±

1.2

− 36.8

±

1.1

179

Mature

− 29.9

±

0.8

− 33.5

±

1.3

− 34.1

±

1.0

Mature

− 31.1

±

0.4

− 35.9

±

0.7

− 34.3

±

0.7

Mature

− 30.1

±

0.5

− 36.3

±

1.0

− 37.0

±

1.1

195

Mature

− 30.1

±

0.9

− 34.2

±

1.7

− 34.9

±

1.4

Mature

− 31.4

±

0.7

− 36.2

±

0.8

− 34.6

±

1.0

Mature

− 30.3

±

0.6

− 36.5

±

1.2

− 37.2

±

1.7

216

Mature

− 29.8

±

0.6

− 33.9

±

0.9

− 34.9

±

0.6

Mature

− 31.2

±

0.1

− 35.8

±

0.3

− 34.4

±

0.2

Mature

− 30.2

±

0.6

− 36.7

±

0.9

− 37.4

±

0.9

239

Mature

− 29.6

±

0.7

− 33.9

±

1.1

− 35.0

±

1.2

Mature

− 31.1

±

0.2

− 35.3

±

0.6

− 34.0

±

0.1

Mature

− 29.8

±

1.1

− 36.9

±

0.5

− 37.6

±

0.7

250

Mature

− 29.3

±

1.2

− 33.5

±

1.9

− 34.7

±

1.3

Mature

− 31.1

±

0.7

− 35.2

±

1.0

− 33.8

±

0.9

Mature

− 29.8

±

1.3

− 37.5

±

0.8

− 38.0

±

0.8

274

Mature

− 29.5

±

0.6

− 33.9

±

0.8

− 35.1

±

0.4

Mature

− 31.8

±

0.8

− 36.1

±

0.6

− 34.7

±

0.6

Mature

− 30.3

±

0.8

− 36.8

±

0.7

− 36.9

±

0.8

Fig. 4

Carbon isotope values of bulk leaf and n-alkanes from a Betula occidentalis, b Populus angustifolia, and c Acer negundo. Carbon isotope values of bulk leaf materials are shown in closed diamonds, while data from n-C27, n-C29, and n-C31 are shown in closed, open, and crossed squares, respectively. Error bars represent the standard deviations around the mean of the three individuals from each species used in this study. Colored areas correspond to the bud (white), leaf expansion (dark gray), and mature leaf (light gray) phenophases

The carbon isotope values of prominent n-alkane homologs are reported in Table 4 and Fig. 4. The δ13C of n-C27 and n-C29 was measured in all samples from three individuals of both B. occidentalis and P. angustifolia, with n-C27 and n-C29 the most abundant homolog for either species, respectively. The δ13C of n-C29 and n-C31 was measured in all samples from three individuals of A. negundo, with n-C31 being the most abundant homolog (Table S2). We found that the δ13C of n-C27 and n-C29 from B. occidentalis and P. angustifolia was highly correlated (R2 = 0.74, p < 0.0001; R2 = 0.60, p = 0.0006, respectively), as was the δ13C of n-C29 and n-C31 from A. negundo (R2 = 0.99, p < 0.0001). Here, we discuss the carbon isotope values of only the most abundant homolog for each species (δ13Cn-alkane) and, thus, we focus our description and discussion on δ13C of n-C27, n-C29, and n-C31 for B. occidentalis, P. angustifolia, and A. negundo, respectively.

As with δ2Hn-alkane, the δ13Cn-alkane varied systematically during specific phenological intervals in each species. Where the patterns in δ2Hn-alkane and δ13Cbulk were generally similar between the species, the patterns in δ13Cn-alkane were relatively unique for each species in this study. During the bud phenophase and prior to leaf flush, the δ13Cn-alkane of both B. occidentalis and P. angustifolia became more 13C-depleted (− 1.2 and − 1.7‰, respectively), whereas the δ13Cn-alkane of A. negundo became more 13C-enriched by 0.6‰. During leaf expansion, the δ13Cn-alkane of both B. occidentalis and A. negundo continued to become 13C-depleted by − 1.3 and − 4.2‰, respectively. During this interval, the δ13Cn-alkane of P. angustifolia abruptly became more 13C-enriched by 2.1‰. In the mature leaf phenophase, δ13Cn-alkane values in all species were steady, where the mean δ13Cn-alkane values were − 33.8 ± 0.3‰ (1σ, n = 6), − 34.3 ± 0.3‰ (1σ, n = 7), and − 37.3 ± 0.4‰ (1σ, n = 6) for B. occidentalis, P. angustifolia, and A. negundo, respectively.

The apparent carbon isotope fractionation (εC) between δ13Cn-alkane and δ13Cbulk is shown in Fig. 5. As with εH, εC varied systematically across the growing season for all species. In general, we found that the εC values became relatively more negative during the bud phenophase for B. occidentalis and P. angustifolia, while εC values became relatively less negative during the bud phenophase in A. negundo. Following leaf flush, the εC values of B. occidentalis initially continued to become less negative throughout leaf expansion phenophase (DOY 141 to 167). Following leaf flush and during leaf expansion, εC values from P. angustifolia immediately became less negative (DOY 146 to 161), while εC from A. negundo transitioned to more negative values during the same phenological stages. Following leaf expansion, the εH values in all species become relatively stable, corresponding to limited variation in both δ13Cn-alkane and δ13Cbulk values. In the mature leaves, the mean εC was − 5.2 ± 0.5‰ (1σ, n = 6), − 3.2 ± 0.3‰ (1σ, n = 7), and − 7.5 ± 0.6‰ (1σ, n = 6) for B. occidentalis, P. angustifolia, and A. negundo, respectively.
Fig. 5

Apparent fractionation between the carbon isotope values of bulk leaf material and n-alkanes from Betula occidentalis, Populus angustifolia, and Acer negundo during the studied period

Discussion

Seasonal dynamics of n-alkane concentrations and distributions

Leaf wax n-alkanes and other lipids are generally considered an adaptation to reduce water loss from the leaf, to provide a photoprotective layer, and to deter pathogens (Eglinton and Hamilton 1967; Guiz 1994; Riederer and Markstadter 1996; Shepherd and Griffiths 2006). The concentrations of n-alkanes can be shaped by a plant’s environment (Bondada et al. 1996; Riederer and Schneider 1990; Shepherd and Griffiths 2006) with the mean concentration of 506 ± 497 µg g−1 observed in angiosperm leaf material (Bush and McInerney 2013). Here, we found the mean concentration of n-alkanes to be 486 ± 510, 825 ± 342, 1064 ± 859 µg g−1 for B. occidentalis, P. angustifolia, and A. negundo, respectively (Table 1). The mean concentrations of n-alkanes for B. occidentalis were not statistically different from the average angiosperm, while both n-alkane concentrations for P. angustifolia, and A. negundo were significantly higher from the average angiosperm (t-test, p = 0.0147 and p < 0.0001, respectively). Nonetheless, concentrations of total and individual n-alkanes were not constant and varied throughout the study period. In general, the concentration of total n-alkanes dropped precipitously from the initial collection in the bud phenophase, likely due to increased mass from bud swelling (Fig. 1). During the leaf expansion phenophase, the concentration of total n-alkanes stabilized or slightly increased, suggesting lipid synthesis and production during leaf expansion. Following leaf expansion, the concentration of total n-alkanes stabilized or decreased in all species, pointing toward either decreased n-alkanes production, leaf wax ablation, or increased leaf mass from leaf thickening. We cannot distinguish among these possibilities, but note that similar seasonal dynamics in the concentration of total and individual n-alkanes has been previously observed in studies of broad-leaf deciduous angiosperms (Jetter and Schaffer 2001; Newberry et al. 2015; Oakes and Hren 2016; Piasentier et al. 2000; Sachse et al. 2009). After leaf flush and the initial leaf lipid formation, variations in leaf surface area likely have a significant impact on the concentration of n-alkanes resulting in an apparent decrease in lipid concentration (Freimuth et al. 2017). Here, a finite amount of n-alkane and other lipids likely were distributed across a broadening and thickening leaf, resulting in an apparent decrease in n-alkane concentration. Thus, it was difficult to assess if the decreases in the concentration of n-alkanes were a result of ablation or changes in leaf morphology (Fig. 1). Recently, Freimuth et al. (2017) related concentration to leaf area, rather than dry mass, and found that the apparent decrease in concentration was likely due to dilution of initially produced n-alkanes over an expanding leaf area. Thus, these data suggest that the apparent seasonal variations of n-alkane concentration are both a function of the initial synthesis of these compounds and changes in leaf morphology.

Odd-over-even distribution of n-alkanes is a characteristic of higher plant n-alkanes (Eglinton and Hamilton 1967). The distribution of n-alkanes from B. occidentalis, P. angustifolia, and A. negundo was broadly consistent with previous observations of the carbon preference indices (CPI) of n-alkanes (Bush and McInerney 2013). The mean CPI of angiosperm n-alkanes is 11.8 ± 12.3 (Bush and McInerney 2013). Here, we found the overall mean CPI of 21.8 ± 12.1, which was significantly different from that of an average angiosperm (t-test, p < 0.0001). Furthermore, we found significant variations in CPI relative to phenophases within the individual species (Table 1). Individually, the mean CPI for A. negundo was not statistically different from the average angiosperm; however, the mean CPI for B. occidentalis and P. angustifolia was significantly different from the average angiosperm (t test, p < 0.0001). The mean CPI of the mature leaf from B. occidentalis was 20.4 ± 3.5 and significantly different from the CPI of the bud and leaf expansion phenophases (37.9 ± 8.5 and 30.8 ± 9.8, respectively; t test, p < 0.0001). Furthermore, the mean CPI for each phenophase of A. negundo was significantly different from each other (bud: 4.6 ± 0.8, leaf expansion: 5.9 ± 0.7, mature leaf: 11.3 ± 1.3), while the mean CPI for each phenophase of P. angustifolia was not different from each other. CPI is an indication of the odd-over-even character of a species’ n-alkane distribution. We observed significant variations across the growing season in two species, one of which became more strongly weighted toward odd chain lengths (e.g., A. negundo) and the other became less strongly weighted toward odd chain lengths (e.g., B. occidentalis). The distribution of n-alkanes on leaves has been linked to various environmental parameters (Bush and McInerney 2013; Shepherd and Griffiths 2006); however, the reasons for the removal or production of specifically odd or even n-alkane chain lengths following the initial formation remain abstruse.

The average chain length (ACL) of leaf waxes is a commonly applied parameter to assess the timing and variations in leaf wax development. Here, we noted systematic variations in ACL across the season corresponding to specific leaf developmental phases in all species, where the ACL values increased during leaf expansion and then stabilized (Fig. 1). Similar seasonal patterns in ACL have been observed in studies of other temperate angiosperm species (Freimuth et al. 2017; Newberry et al. 2015; Oakes and Hren 2016; Sachse et al. 2015; Tipple et al. 2013). These studies have interpreted this pattern to reflect the vast production and cessation of leaf wax development during the early ontogeny of the leaf. However, we noted that the ACL of both P. angustifolia and A. negundo stabilized following leaf expansion phenophase, while B. occidentalis demonstrates a more protracted transition towards stabilization during the mature leaf phenophase in the mid-summer. We observed that the three B. occidentalis specimens had greater variation in ACL between the individual specimens compared to P. angustifolia and A. negundo (Table S1). Studies of other angiosperm species have shown relationships between n-alkane distributions and environmental temperature, where higher growth temperatures were correlated with increased ACL (Bush and McInerney 2015; Maffei et al. 1993; Riederer and Schneider 1990; Sachse et al. 2006; Tipple and Pagani 2013). ACL from a variety of angiosperm species has also corresponded to plant–water relations, where drier, more water-limited conditions were correlated with increased ACL (Hughen et al. 2004; Rommerskirchen et al. 2003, 2006; Schefuss et al. 2003; Shepherd et al. 1995; Vogts et al. 2009, 2012). The distribution and composition of n-alkanes and other leaf waxes from specific species have been shown to be relatively plastic and adaptable to a degree, controlled largely by the plant’s environmental conditions (Shepherd and Griffiths 2006). However, it is unclear how much variability in ACL could occur among individuals of the same species, growing in similar locations and environments. One possible explanation for the protracted transition to stability is that individual leaves may be experiencing slightly different microclimates. If this were the case, we would expect relatively high variability throughout the growing season. However, the ACL of the three individuals of B. occidentalis converged toward the end of the growing season (Fig. 1). Additionally, it may be possible that individual plants of the same species have slightly different leaf ontogenies, with specific individual plants lagging or preceding others. Further, the protracted transition may be the result of continued leaf wax production following the initial formation. However, in this scenario we would expect the δ2H and δ 13C values to possibly vary in response. We will explore this hypothesis more in Sects. Interspecies differences and similarities in the δ2H of plant waters and n-alkanes and Temporal trends δ13C of n-alkanes and possible mechanisms. Nonetheless, unlike concentrations of n-alkanes relative to dry weight, ACL is independent of changes in leaf mass and the relatively constant ACL after leaf maturity suggests that leaf wax production largely ceases in mature leaves of broad-leaf angiosperm trees.

Interspecies differences and similarities in the δ 2H of plant waters and n-alkanes

Major research areas regarding the δ2H values of leaf waxes have focused on (a) establishing the relationship between the δ2H values of n-alkanes and precipitation, (b) understanding when the δ2H signals in waxes are established, and (c) determining what are the factors controlling the δ2H values of lipids during leaf and lipid development. Understanding the relationships between the δ2H values of n-alkanes and precipitation and the timing of leaf wax formation in different plant functional types, leaf morphologies, and growth habits have been recognized as significant for geologic applications (e.g., Diefendorf and Freimuth 2017; Sachse et al. 2012). These factors, as well as, the aerial coverage, productivity, and relative concentration of leaf lipids of certain species or plant types, are important in developing robust paleo-reconstructions of plant community dynamics and hydrologic cycle variations with the isotope values of leaf wax lipids (Tipple et al. 2013). Carefully controlled natural and laboratory experiments are necessary to develop an understanding of the biological and environmental factors governing the δ2H values of leaf waxes (Kahmen et al. 2011, 2013b; Oakes and Hren 2016; Tipple et al. 2015). These studies either control or monitor the δ2H values of environmental water sources, as well as the δ2H values of n-alkanes. As with all hydrogen within a plant’s tissue, the hydrogen in n-alkanes is largely derived from the plant’s source water and, thus, the δ2H values of source water. The δ2H value of source water can be altered after it enters the plant, and as the initial biosynthesis of n-alkanes occurs within the leaf chloroplast, the water used for synthesis can be altered due to leaf water evaporation as well as isotopic exchange with atmospheric water vapor. Several natural and controlled growth experiments have shown strong relationships between δ2H value of leaf water and n-alkanes indicating that leaf water 2H-enrichment is transferred to n-alkanes (Kahmen et al. 2013a, b; Tipple et al. 2015). The fractionation between biosynthetic water associated with n-alkane and other leaf wax lipid formation remains enigmatic and an active area of research. Biosynthetic water cannot be measured; however, leaf water is related to biosynthetic water and the δ2H value of leaf water has often been used to estimate the biosynthetic fractionation (Freimuth et al. 2017; Gamarra et al. 2016; Kahmen et al. 2013a, b; Newberry et al. 2015; Tipple et al. 2013, 2015).

Our study site, Big Cottonwood Canyon, Utah, was specifically identified as an ideal location to conduct a semi-controlled, natural experiment where the biological and environmental factors controlling the δ2H values of n-alkane (δ2Hn-alkane) could be isolated. At this site, the δ2H values of stream water (δ2HMW), and most likely source water, were relatively stable throughout the growing season (Table 2), eliminating one of the main variables that control the δ2H values of plant waters and n-alkanes. Plants take up soil water with no isotopic fractionation (Dawson and Ehleringer 1993) and we found that the δ2HXW values from all three species were very similar to the δ2HMW values after the cessation of leaf flush (Table 2). Nonetheless, we found that the δ2H values of stem water (δ2HXW) diverged from the δ2HMW during the bud and leaf expansion phenophases. During these phases, the δ2HXW of all three species was less negative than the δ2HMW. Similar patterns in the δ2HXW have been observed in other broad-leaf angiosperm trees (Freimuth et al. 2017; Martin-Gomez et al. 2017; Phillips and Ehleringer 1995; Sachse et al. 2015) and have been argued to be the signature of residual evaporatively enriched xylem water from the previous winter (Phillips and Ehleringer 1995). The trend in δ2HLW corresponded to patterns in the δ2HXW during the bud and leaf expansion phenophases (Fig. 2). Following leaf expansion, the δ2HLW quickly diverged from the δ2HXW and became much less negative. The probable reason for this offset and sudden divergence is that the leaf water was affected by both evaporation and exchange with atmospheric water vapor and once the leaf was fully expanded the impact of processes was amplified. The dynamic relationship between the δ2HLW, δ2H values of the atmospheric water vapor, and climate is well established (e.g., Dawson et al. 2002). Nonetheless, the fixed δ2H values of the stream and xylem water after leaf expansion allow the effects of leaf water 2H-enrichment to be highlighted.

The δ2Hn-alkane from all three species also demonstrated isotopic variations corresponding to leaf developmental phenophase. We found that δ2Hn-alkane from all species was lowest in the bud phenophase, became relatively 2H-enriched throughout the leaf expansion phenophase, and stabilized after leaf expansion. These data are consistent with several previously observations of changes in δ2Hn-alkane across a growing season and have been explained as the rapid de novo production of leaf waxes during the leaf flush and expansion phenophase followed by cessation of leaf wax development once the leaf was fully expanded and mature (Freimuth et al. 2017; Oakes and Hren 2016; Sachse et al. 2015; Tipple et al. 2013). Here, we found that δ2Hn-alkane during the mature leaf phases was statistically different (t test, p ≤ 0.0096) from one another. While both B. occidentalis and A. negundo had similar leaf phenology—leafing out during the same interval—the δ2Hn-alkane from mature leaves was different. These interspecies differences could be due to different strategies for de novo synthesis, variations in H-sources, or possibly unique biosynthetic fractionations. We observed differences in the temporal expression of the δ2H values of n-alkanes from B. occidentalis and A. negundo, with the δ2Hn-alkane stabilizing slightly after leaf expansion in B. occidentalis, whereas in A. negundo, the δ2Hn-alkane is stabilized at the initial leaf expansion (Fig. 2). These differences may possibly be due to species-specific strategies for de novo synthesis of leaf waxes. One possible mechanism may relate to differences in ecology, where A. negundo are much more cosmopolitan, thriving in many environments relative to the exclusively riparian B. occidentalis. A. negundo is adapted to a wider variety of conditions, including more water-stressed environments and, as a result, A. negundo may develop its cuticular leaf waxes much earlier in the growing season to preempt possible variations in early season conditions. Nonetheless, the temporal expression of the δ2Hn-alkane from P. angustifolia is similar to A. negundo, and P. angustifolia is exclusively a riparian species. These patterns may also be a response to differences in growth form, where both P. angustifolia and A. negundo are fast growing, upright tree species, while B. occidentalis presents a more multi-stemmed form. The δ2Hn-alkane from multi-stemmed shrubs has been shown to be relatively less negative than tree species (Sachse et al. 2012)—consistent with our findings here. This pattern in shrubs has been reasoned to reflect leaf water enrichment associated with more xeric shrub habitats (Sachse et al. 2012). Although here, the site location was riparian (mesic) and the source water and environmental conditions were identical for these three species; thus, the differences in absolute δ2Hn-alkane between these species must be related to additional factors.

The apparent biosynthetic fractionation between the δ2H values of leaf water and n-alkane (εH) in all species exhibited significant variations throughout the growing season. εH was least negative during or immediately following the leaf expansion phenophase (Fig. 3) and was similar to those previously reported during leaf expansion (Freimuth et al. 2017; Newberry et al. 2015; Sachse et al. 2015). However, during the mature leaf phenophase, we observed εH values that became progressively more negative as the season progressed (Fig. 3). The overall mean εH during this period was − 148 ± 27‰ (1σ, n = 13). By tracing both the δ2Hn-alkane and δ2HLW, it was evident that this variation was due to enrichment of 2H in the leaf water and not changes in the δ2Hn-alkane value (Fig. 2). These data suggest that in natural systems, εH is only meaningful during the time of leaf wax development and will fluctuate post-development due to changes in the δ2HLW of leaf water. While determining the “true” biosynthetic fractionation is not possible, greenhouse studies where the δ2H of source water and atmospheric water vapor have been maintained throughout leaf wax develop may be the best approximation. Greenhouse studies using B. occidentalis and a similar Populus species, P. fremontii, found εH of − 150 ± 9 and − 164 ± 10‰ for these species, respectively (Tipple et al. 2015). While both this and the greenhouse study demonstrate variability in εH, the mean εH values calculated in this study were less negative and variable, suggesting a contribution of more 2H-enriched H-source in the field-grown plants. One difference between this study and the greenhouse study was that this study used mature trees and the greenhouse study used saplings. Potentially, the mature trees were able to utilize stored 2H-enriched carbohydrates in leaf wax lipid production, whereas the saplings were more dependent of recent photosynthate for lipid formation. Nonetheless, this suggests that “true” biosynthetic fractionation is largely related on the δ2HLW during the period of de novo n-alkane synthesis and aberrations in εH in broad-leaf angiosperms are related to variations in δ2HLW after leaf wax formation.

Temporal trends δ 13C of n-alkanes and possible mechanisms

Carbon isotope values of plant tissues are used to track the transition from heterotrophic to autotrophic growth in plants and to assess offsets between photosynthetic and non-photosynthetic plant tissues (Cernusak et al. 2009; Dietze et al. 2014; Kimak et al. 2015; Leavitt and Long 1982, 1985; Li et al. 2007; Terwilliger 1997; Terwilliger et al. 2001). Within C3 plants, non-photosynthetic stems, roots, fruits, and seeds, as well as emerging leaves tend to be 13C-enriched relative to fully mature leaf tissue (Brugnoli and Farquhar 2000; Cernusak et al. 2002, 2007; Park and Epstein 1960). Given the importance of non-photosynthetic plant materials in paleoclimate reconstructions, such as tree ring cellulose and leaf wax n-alkanes, it remains a challenge to develop mechanistic understandings of these carbon isotopic offsets. However, a mechanistic understanding of the relationship between the δ13C of n-alkanes and leaf tissues remains imperfect. Previous analyses of the δ13C of n-alkanes across a growth period showed either little to no change or significant variations in δ13Cn-alkane (Pedentchouk et al. 2008). Nonetheless, it is difficult to determine the underlying mechanisms for these variations as leaf developmental phenophase was not noted. Here, we compared the δ13C of n-alkanes and leaf tissues across the growing season in relation to leaf development to assess the heterotrophic–autotrophic transition between these two materials.

Deciduous species remobilize reserve carbohydrates from previous years to initially build their photosynthetic tissues. These carbohydrates are converted into metabolic intermediates and utilized in the biosynthesis of other compounds and tissues. The δ13C of emerging leaves tends to become initially less negative and then became more negative as the new leaf became photosynthetically competent. The reason for this δ13C pattern in new leaves remains a subject of debate, but developmental variations in photosynthetic discrimination against 13C may be a possible explanation (Cernusak et al. 2009). We found that the δ13C of bulk leaf material (δ13Cbulk) was consistent with these previous studies (Fig. 4). Here, the δ13Cbulk of these three species became less negative prior to leaf flush and then became more negative until the leaf was fully expanded. One hypothesis that has been put forward to explain the 13C-enrichment followed by 13C-depletion in young leaves is that these leaves undergo a shift in the discrimination related to leaf ontogeny, where the expanding leaves have a greater discrimination against 13C than mature leaves and/or the re-fixation of respired CO2 before young leaves are capable of net CO2 fixation (Cernusak et al. 2001, 2009; Francey et al. 1985; Terwilliger 1997; Terwilliger et al. 2001).

Where the bulk leaf material from all three species showed the expected δ13C pattern from remobilization and incorporation of stored carbon into the leaf tissue, the pattern in the δ13C of n-alkanes (δ13Cn-alkane) varied and was not consistent between the three species. We found that each species had a unique pattern in δ13Cn-alkane during the bud and leaf expansion phenophases, suggesting the possible differences in carbon metabolism and/or biosynthesis of n-alkanes during the initial development of leaf waxes. The δ13Cn-alkane from A. negundo showed a slight enrichment in 13C during the bud phenophase (i.e., < 1‰), followed by a ~ 4‰ depletion in 13C during leaf expansion (Fig. 4). While this pattern broadly mirrors that observed in the δ13Cbulk, the absolute values were different. This pattern is consistent with the current understanding of remobilization and incorporation of stored carbon from previous growth season into young leaf tissues and non-photosynthetic components, followed by the incorporation of recently fixed carbon into these materials once the leaf becomes photosynthetically competent. Similar variations across the growing season have also been observed in the δ13C values of leaf cellulose of several angiosperm tree species (Kimak et al. 2015). On the other hand, both the B. occidentalis and P. angustifolia demonstrated a 1-2‰ depletion in 13C during the bud phenophase, followed by a ~ 1–2‰ enrichment in 13C during the leaf expansion phenophase (Fig. 4). This pattern indicates that fatty acid synthesis and elongation are decoupled from photosynthetic tissue development during this period of development. This pattern may also suggest that the carbon stock used in the production of n-alkyl leaf wax lipids during the early growing season may derive from a different store relative to other leaf tissues or additional isotopic branch points may exist in some species (Hayes 1993). Nonetheless, the comparison of the δ13Cn-alkane of these three species suggests that strategies of leaf wax development may not be consistent between broad-leaf angiosperm tree species.

The apparent fractionation between the δ13C values of leaf material and n-alkane (εC) in each species demonstrated significant variations related to leaf development phenophase. We observed large (i.e., > 3‰) variations in εC in all species during the initial growing season followed by relative stability after the leaf fully expands (Fig. 5). As with δ13Cn-alkane, the patterns in εC suggest a decoupling between leaf development and leaf wax formation. The transient nature of the εC signal may suggest the isotopic variations in both the leaf material and n-alkane due to variations in the stored carbon materials utilized during the initial growing season. Generally, the εC is calculated from the δ13C values of leaf material and n-alkanes on fully mature leaves. The mean εC between δ13C values of leaf material and n-C27, n-C29, and n-C31 alkanes for temperate angiosperm tree species is − 4.6 ± 2.0‰ (Diefendorf and Freimuth 2017). Here, we found that the mean εC for mature leaves from the three species was unique from each other (t test, p < 0.0001), but only the εC from A. negundo was unique from the mean εC from temperate angiosperm tree species (t test, p = 0.0006). The εC value has been shown to vary among closely related tree species grown in the same environment (Diefendorf et al. 2015b). In particular, the εC of numerous conifer species has been related to the Family level, with these distinctions hypothesized to be related to differences in carbon storage, allocation, and metabolism (Diefendorf et al. 2015b; Freeman and Pancost 2013). While the biochemical mechanisms underlying these patterns are unknown, these data suggest that coupling the carbon isotope values of leaf materials and leaf wax lipids in controlled growth environments may be a productive method to uncover the fundamental mechanisms driving these isotopic differences and the ecological information encoded within these isotopic signals.

Coupling temporal patterns of hydrogen and carbon isotope values of n-alkanes

As described above, the δ2H and δ13C values of n-alkanes individually provide information on plant–water relations and growth environment, respectively. When both are measured in the same material, a significant relationship has been observed between δ2H and δ13C values of n-alkanes and other n-alkyl lipids within individual plants and plants using the same photosynthetic pathway (Chikaraishi and Naraoka 2003; Chikaraishi et al. 2004b; Feakins and Sessions 2010b; Pedentchouk et al. 2008). The apparent fractionations, εH and εC, have also been shown to be distinctive between photosynthetic pathways and have been suggested as a method to distinguish potential sources of n-alkanes and other lipids in geologic materials (Chikaraishi and Naraoka 2003; Chikaraishi et al. 2004b). These relationships have been used to interpret the δ2Hn-alkane and δ13Cn-alkane from ancient plants (Magill et al. 2013; Tipple and Pagani 2010). However, the seasonal variations in and between δ2Hn-alkane and δ13Cn-alkane are relatively unknown.

The relationship between δ2Hn-alkane and δ13Cn-alkane varied with leaf development phenophase (Fig. 6). We found relatively consistent trends in δ2Hn-alkane and δ13Cn-alkane within the bud, leaf expansion, and mature leaf phenophases. A linear trajectory was observed between the δ2Hn-alkane and δ13Cn-alkane within the bud phenophase; however, A. negundo demonstrated a slight positive correspondence, while B. occidentalis and P. angustifolia showed negative correspondences. The δ2Hn-alkane and δ13Cn-alkane from the leaf expansion phenophase were transitory, characterized by values changing towards the values of the mature leaf phenophase (Fig. 6). As expected, the δ2Hn-alkane and δ13Cn-alkane from the mature leaf phenophase were relatively constant throughout the remainder of the growing season, with all three species occupying unique δ2Hn-alkane/δ13Cn-alkane space.
Fig. 6

Cross-plot of the carbon and hydrogen isotope values of n-alkanes during the a bud, b leaf expansion, and c mature leaf phases. Data from Betula occidentalis, Populus angustifolia, and Acer negundo shown in open, gray, and closed circles, respectively

The transition from heterotrophy to autotrophy within leaf wax production could be identified when δ2Hn-alkane and δ13Cn-alkane were coupled and monitored throughout the growing season. Here, we observed that the initial δ2Hn-alkane and δ13Cn-alkane values from buds reflected residual waxes, likely produced during the previous growing season. Once the bud became active, new n-alkanes were produced with a hydrogen source material that was 2H-enriched relative to residual bud lipids. The carbon source material utilized to produce n-alkanes during this heterotrophic phase appeared to be either 13C-enriched or -depleted depending on species. These data suggested that the precursors to the NADPH and pyruvate utilized in initial production of leaf wax n-alkanes might be variable. It remains unclear if these signals change annually or if these species continually utilize similar precursors. Once the leaf became photosynthetic active, relatively large variations in the δ2Hn-alkane and δ13Cn-alkane were apparent, suggesting that new photosynthate and biosynthetic water was utilized during this later leaf expansion period. Following leaf maturity, the δ2Hn-alkane and δ13Cn-alkane were unchanged indicating little to no new leaf wax production. Thus, the final δ2Hn-alkane and δ13Cn-alkane values represented a mixture of isotopic signatures from the heterotrophic and autotrophic phases of leaf wax development.

Notes

Acknowledgements

This research was supported by NSF-ISO-1052551. The authors thank Bastian Hambach, Christine Doman, and Susanna Khachaturyan for their assistance in the analysis of the carbon isotope values of leaf waxes, hydrogen isotope values of leaf waxes, and hydrogen isotope ratios of plant and environmental waters, respectively. The authors also wish to thank Katherine H. Freeman and Ansgar Kahmen for helpful insights and discussions that strengthened this contribution.

Author contribution statement

BJT and JRE conceived and designed experiment. BJT performed the experiment and analyzed the data. BJT and JRE wrote the manuscript.

Supplementary material

442_2018_4189_MOESM1_ESM.xlsx (71 kb)
Supplementary material 1 (XLSX 70 kb)
442_2018_4189_MOESM2_ESM.xlsx (48 kb)
Supplementary material 2 (XLSX 47 kb)

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© Springer-Verlag GmbH Germany, part of Springer Nature 2018

Authors and Affiliations

  1. 1.Department of BiologyUniversity of UtahSalt Lake CityUSA
  2. 2.Global Change and Sustainability CenterUniversity of UtahSalt Lake CityUSA

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