# Cuticle thickness affects dynamics of volatile emission from petunia flowers

Pan Liao, Shaunak Ray, Benoît Boachon, Joseph H Lynch, Arnav Deshpande, Scott Mcadam, John A Morgan, Natalia Dudareva

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Pan Liao, Shaunak Ray, Benoît Boachon, Joseph H Lynch, Arnav Deshpande, et al. Cuticle thickness affects dynamics of volatile emission from petunia flowers. Nature Chemical Biology, 2020, 17 (2), pp.138 - 145. 10.1038/s41589-020-00670-w. hal-04701247

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Cuticle thickness affects dynamics of volatile emission from petunia flowers

Pan Liao¹,#, Shaunak Ray²,#, Benoît Boachon¹,†, Joseph H. Lynch¹,³, Arnav Deshpande²,
Scott McAdam³,⁴, John A. Morgan¹,², Natalia Dudareva¹,³,⁵,*

¹Department of Biochemistry, Purdue University, West Lafayette, IN 47907-2063, USA;

²Davidson School of Chemical Engineering, Purdue University, West Lafayette, IN 47907-2100, USA;

³Purdue Center for Plant Biology, Purdue University, West Lafayette, IN 47907, USA.

⁴Department of Botany and Plant Pathology, Purdue University, West Lafayette, IN 47907, USA.

⁵Department of Horticulture and Landscape Architecture, Purdue University, 625 Agriculture Mall Dr., West Lafayette, IN 47907-2010, USA.

#Equal contributions

*Correspondence: dudareva@purdue.edu

†Present address: BVpam FRE 3727, Université de Lyon, Université Jean Monnet Saint-Etienne, CNRS, Saint-Etienne, France.
**Editorial Summary:** The plant cuticle is not simply a passive diffusion barrier that volatiles cross before release to the atmosphere, but also a sink/concentrator for volatiles, which protects cells from toxic effects of these hydrophobic compounds.

**Abstract**

The plant cuticle is the final barrier for volatile organic compounds (VOCs) to cross for release to the atmosphere, yet its role in the emission process is poorly understood. Here, using a combination of reverse-genetic and chemical approaches, we demonstrate that the cuticle imposes significant resistance to VOC mass transfer, acting as a sink/concentrator for VOCs and hence protecting cells from the potentially toxic internal accumulation of these hydrophobic compounds. Reduction in cuticle thickness has differential effects on individual VOCs depending on their volatility, and leads to their internal cellular redistribution, a shift in mass transfer resistance sources and altered VOC synthesis. These results reveal that the cuticle is not simply a passive diffusion barrier for VOCs to cross, but plays the aforementioned complex roles in the emission process as an integral member of the overall VOC network.
**Introduction**

Plants emit annually over 10⁹ metric tons of volatile organic compounds (VOCs), thus significantly contributing to atmospheric chemistry and climate¹,². Released from flowers, fruits and vegetative tissues, VOCs play critical roles in plant fitness by attracting pollinators and other beneficial organisms, defending plants against herbivores and pathogens, facilitating plant-plant communication, and protecting against abiotic stresses³,⁴. Due to the plethora of biological processes dependent on VOCs, chemical composition and quantity of emitted volatiles are species-, organ- and tissue-specific, developmentally and/or temporally regulated, and depend on biotic and abiotic factors³,⁵⁻⁷. Significant progress has been made in understanding the biosynthesis of plant VOCs and their regulation⁴,⁸, but much less is known about how volatiles are transported from cells into the atmosphere.

Although chemically diverse, VOCs are lipophilic low-molecular-weight (~100-200 Da) compounds with high vapor pressure at ambient temperature. To be released from cells, they must move from the site of synthesis through the cytosol, cross the plasma membrane, hydrophilic cell wall, and finally the waxy cuticle⁹. Until recently, it was assumed that VOCs simply diffuse through each barrier. However, modeling of emission predicted that VOCs would accumulate to toxic levels in membranes in the absence of biological mediation due to their favorable partitioning into the lipid bilayers⁹. These *in silico* model predictions were experimentally validated *in planta* by down-regulation of an ATP-binding cassette (ABC) transporter, *PhABCG1*, capable of transporting phenylpropanoid/benzenoid volatiles across the plasma membrane in petunia flowers¹⁰.

The cuticle is the final physical barrier for VOCs to cross in most plant cells. Due to their hydrophobicity, the rate of VOC transport should depend on the physicochemical properties of
the compound itself (diffusivity and solubility in the cuticle) and the cuticle composition and thickness<sup>11-14</sup>. Although cuticle structure and composition vary considerably between plants, organs and developmental stages, it is commonly composed of cutin and cuticular wax, the latter of which is embedded in and deposited on the surface of the cutin matrix<sup>12,15</sup>. The cuticular wax is a semi-crystalline mixture of alkanes, aldehydes, primary and secondary alcohols, ketones and esters, all derived from very-long-chain fatty acids<sup>14,16,17</sup>. The cuticle generally acts as a molecular transport barrier, protecting tissues from stresses and preventing organ fusion during plant development<sup>15</sup>. While the role of the cuticle in transpirational water flux is well established<sup>15,18</sup>, and extensive prior research provides knowledge about the interaction of atmospheric VOCs with the cuticle<sup>19-23</sup>, how the cuticle is involved in the release of hydrophobic volatiles into the atmosphere remains an unresolved question.

Here, we use *Petunia hybrida* flowers, which emit high levels of phenylalanine (Phe) derived volatiles<sup>24,25</sup>, to investigate the role of the cuticle in the emission process. We show that reduction in cuticle thickness not only alters VOC emission, but also leads to redistribution of VOC internal pools and feedback inhibition of VOC biosynthesis, demonstrating that cuticle is an integral member of the overall emission network.

## Results

**Generation of petunia flowers with altered cuticles.** Our mathematical modeling of VOC emission predicted that out of all barriers the cuticle imposes the highest resistance to VOC efflux<sup>9</sup>, yet this has not been experimentally evaluated. To determine whether cuticle of petunia flowers retains VOCs, their levels were analyzed in cuticular waxes of flowers on day 2
postanthesis, the developmental stage with the highest VOC emission²⁶. This analysis revealed that cuticle holds 53 ± 4% of total VOC internal pools (Fig. 1a).

Previous studies have shown that cuticle formation can be altered via perturbing transport of wax precursors from the site of their biosynthesis at the endoplasmic reticulum across the plasma membrane and through cell wall to be deposited at the cell surface²⁷. It is well established that transport of cuticular monomers across the plasma membrane requires ABC transporters¹⁵. In Arabidopsis, loss-of-function mutants of any of three closely related transporters, AtABCG11, AtABCG12, or AtABCG13, result in cuticle-deficient phenotypes²⁸⁻³³. Recently, we identified 17 ABCG transporters expressed in petunia flowers, two of which formed a clade with the three aforementioned Arabidopsis wax transporters¹⁰. These two putative petunia wax transporters, Ph9795 and Ph13519 are designated PhABCG11 and PhABCG12, respectively, as they exhibit 78%/86% and 66%/80% identity/similarity to AtABCG11 and AtABCG12, respectively. In addition, a blastn search against the genome of *P. axillaris*, the parent that mainly contributes to the *P. hybrida* genome³⁴, revealed three and one additional homologs of *AtABCG11* and *AtABCG12*, respectively. However, only *PhABCG11* and *PhABCG12* were expressed in *P. hybrida* corolla at high levels³⁵ (labeled as *PhABCG11a* and *PhABCG12b*, respectively, in Supplementary Tables 1 and 2). Expression of *PhABCG12* was 53% higher than *PhABCG11*, and both genes were developmentally regulated, with highest expression early in flower development, consistent with timing of cuticle formation (Supplementary Figure 1). Neither *PhABCG11* nor *PhABCG12* follow the developmental expression profile of scent biosynthetic genes, which have the highest levels on day 2 postanthesis¹⁰,²⁶. Thus, *PhABCG11* and *PhABCG12* are promising targets for experimental perturbation of cuticle properties for subsequent analysis of the cuticle's role in VOC emission.
Our attempts to downregulate expression of each gene using RNAi approach under the control of a petal-specific promoter ³⁶ resulted in no transgenic lines for *PhABCG11*, while seven lines were obtained for *PhABCG12*. Three lines, *PhABCG12-7*, -8 and -9, with the greatest downregulation (68-79%) were selected for further analysis (Fig. 1b). *PhABCG12-RNAi* construct specifically targeted only *PhABCG12* (Supplementary Figure 2a) without affecting *PhABCG12a* expression (Supplementary Figure 2b). Unexpectedly, *PhABCG11* expression in *PhABCG12-RNAi* lines was coordinately decreased by 36-41% via an unknown mechanism (Supplementary Figure 2c). Total wax loads in *PhABCG12-RNAi* lines were reduced by 35-55% relative to wild-type and empty vector control (Fig. 1c), while chemical composition and ratios between wax constituents were not significantly altered (Extended Data Fig. 1). The flowers of transgenic plants displayed visible changes relative to wild-type and empty vector control. They were smaller, had less smooth surface, reduced fresh weight and more translucent petals (Fig. 2a upper panel and Extended Data Fig. 2). TEM analysis of conical epidermal cells revealed a 22% reduction in cuticle thickness in petals of *PhABCG12-9*, the line with greatest *PhABCG12* downregulation, relative to wild type (Figs. 2b, c, and Extended Data Fig. 3). Toluidine blue (TB) staining, which detects increased cuticle permeability³⁷, showed that petals from transgenic lines, but not wild-type or empty vector control, exhibited Class II staining, characterized by patchy and random pattern³⁷ (Fig. 2a lower panel). Cryo-SEM of transgenic petals revealed that some epidermal cells lost their conical shape and became flattened (Supplementary Figure 3). This morphological change was not uniform across the petal surface, but affected ~23% of the petal area (Supplementary Figure 4), and may result from petal-to-petal fusion prior to flower opening, as was shown in cuticle-deficient mutants³⁰,³². Consistent with reduced cuticle thickness, transgenic *PhABCG12-RNAi*
flowers lost water faster than controls (Supplementary Figure 5). However, phenotypic changes
were not attributable to enhanced water loss, as transgenic plants grown in growth chamber with
higher relative humidity did not recover the wild type size phenotype, VOC emission and their
internal pools (Supplementary Figure 6). Since perturbing the cuticle could have pleiotropic
effects on general cellular functions, we measured levels of total protein, sucrose, starch and
chlorophyll, as well as petal respiration rate, seed yield and germination rate. All parameters
remain unchanged in transgenic flowers relative to controls (Supplementary Figure 7). Overall,
these results demonstrate that altered cuticle thickness is the primary consequence of
*PhABCG12*-RNAi downregulation, enabling analysis of the effect of cuticle on VOC emission.

**Cuticle is an essential member of volatile network.** Translocation of VOCs across the cuticle involves three physical processes: (i) partitioning of VOCs from the aqueous cell wall into the nonpolar waxes, (ii) diffusion across the cuticle, and (iii) vaporization of the VOCs from the cuticle surface. It has been hypothesized that cuticular diffusion is the major barrier to VOC emission⁹, and therefore a thinner cuticle was expected to permit increased VOC release. However, VOC profiling in *PhABCG12*-RNAi flowers revealed the opposite, total emission of VOCs was unexpectedly reduced by 50-56% (Fig. 2d) and their internal pools decreased by 31-43% relative to controls (Fig. 2e). Out of nine individual Phe-derived compounds emitted by petunia flowers, only two low-abundant compounds comprising < 1% of total released VOCs, phenylethylbenzoate and eugenol, remained unaffected by changes in cuticle thickness (Extended Data Fig. 4). The effect of *PhABCG12*-RNAi downregulation on total VOC internal pools was opposite to that previously observed for RNAi downregulation of *PhABCG1*
transporter, which transports VOCs across plasma membrane¹⁰, confirming differential effects
of these two transporters on VOC emission.

Reduction in both emission and internal pools of VOCs in transgenics indicates that VOC
formation is reduced. Indeed, evaluation of the total biosynthetic flux, determined as the sum of
total emission and changes in corresponding internal pools over time, in control and transgenic
flowers showed that biosynthesis was reduced by 52 ± 7.8 % in *PhABCG12-RNAi* flowers (Fig.
2f). To determine where in the scent biosynthetic network inhibition takes place, either upstream
of volatile phenylpropanoid/benzenoid compound synthesis (Phe biosynthesis) or downstream of
Phe (VOC biosynthesis), wild-type and *PhABCG12-9* flowers fed with ¹³C₆-Phe were subjected
to metabolic flux analysis (Extended Data Fig. 5). Similar total biosynthetic fluxes in wild-type
and transgenic flowers after Phe feeding indicate that VOC biosynthetic capacity was not
affected in *PhABCG12-RNAi* flowers (Extended Data Fig. 5c), suggesting that inhibition of
VOC production occurred in the Phe biosynthetic network. This was verified by feeding 2-day-
old wild-type and *PhABCG12-9* flowers with uniformly ¹³C-labeled ([U-¹³C₆]) glucose and
analyzing Phe pool sizes and isotopic abundances. In both genotypes, the glucose labeling
pattern was nearly identical (Supplementary Figure 8a). Over the timecourse, there was little
difference in Phe labeling, despite a 51% reduction in its pool size in transgenic petals at 10 PM
(Supplementary Figure 8b and c), indicating that indeed the carbon flux towards Phe is reduced
in *PhABCG12-9* petals. qRT-PCR analysis of transcript levels of selected genes involved in Phe
biosynthesis revealed that all of them were downregulated by 33 to 62% in *PhABCG12-RNAi*
lines relative to control (Supplementary Figure 9).

While ¹³C₆-Phe feeding compensated for reduced internal pools in transgenics and led to
their similar isotopic labeling in control and transgenic petals (Extended Data Fig. 6a and c), it
did not restore emission in *PhABCG12*-RNAi flowers over 4 h despite its similar to control isotopic labeling (Extended Data Fig. 5d, 6b and d). If the cuticle is the primary resistance to net movement of volatiles out of the cell (mass transfer), it is expected that Phe feeding would result in higher VOC efflux from flowers with thinner cuticle. However, the lag in emission (Extended Data Fig. 6b), which resulted in the inability of transgenic flowers to catch up to wild-type levels of total emission despite rapid internal pool buildup (Extended Data Fig. 6a), suggests the existence of transport limitation(s) upstream of cuticle. Indeed, expression of *PhABCG1*, encoding the plasma membrane-localized VOC transporter, was reduced by 54% in transgenic flowers versus control (Supplementary Figure 9), but total membrane transport capacity remains sufficient to sustain high emission under feeding conditions (Extended Data Fig. 6b).

**Cuticle thickness affects dynamics of individual VOCs.** To account for reduced VOC biosynthesis in comparative analysis of wild type and transgenics, we defined a VOC emission factor (VEF) as the ratio of emission flux to corresponding total biosynthetic flux. A VEF value approaching 1 indicates that VOC emission is controlled mainly by biosynthesis, while VEF values lower than 1 reflect the existence of mass transfer limitations. Contrary to our hypothesis that a thinner cuticle would be more permeable to VOCs, *PhABCG12*-RNAi flowers displayed decreased VEF (Fig. 3a). Comparison of VEF of the individual volatiles in wild type petals revealed that two compounds, methylbenzoate and benzaldehyde, have the highest values close to 1 (Fig. 3b). Out of all emitted compounds, they have the highest volatility based on vapor pressure (Supplementary Table 3) and were more abundant in the mixture of emitted volatiles with relatively small internal pools within the cell (Extended Data Fig. 4). Indeed, emission and biosynthetic fluxes overlaid for each of these two compounds when compared in wild-type
petunia flowers (Fig. 3c), suggesting that methylbenzoate and benzaldehyde emissions are driven primarily by their biosynthesis with the cuticle providing limited resistance for release. In contrast, volatiles with lower ambient vapor pressures such as benzylbenzoate, isoeugenol, 2-phenylethanol, and benzylalcohol (Supplementary Table 3), had lower VEFs (Fig. 3b) and substantially larger internal pools relative to their corresponding emissions (Extended Data Fig. 4). Moreover, these compounds displayed a clear delay between the peak of biosynthesis and emission, and reduction in cuticle thickness did not change these patterns (Fig. 3d, Extended Data Fig. 7), suggesting the existence of a mass transfer limitation for less volatile compounds. Thinner cuticles in transgenics had negligible effect on VEF of methylbenzoate and benzaldehyde, but decreased those of benzylbenzoate, isoeugenol, 2-phenylethanol and benzylalcohol (Fig. 3b). Thus, the trend towards internal build up rather than emission for only less volatile compounds in transgenic petals (Fig. 3b) suggests that effect of cuticle thickness on VOC dynamics depends on physicochemical properties of compounds.

Analysis of VOC internal pools' distribution within the cell revealed that thinner cuticle holds a smaller proportion of VOCs, 32 ± 2% on average in transgenic plants (Fig. 3e) versus 53 ± 4% of total internal pools in wild-type flowers (Fig. 1a). Reduction in cuticle thickness decreased the relative cuticular abundance of all VOCs, although to different degrees, in transgenic petunia flowers relative to control (Extended Data Fig. 8a), leading to their accumulation within the cell. Previously, it was proposed and experimentally confirmed<sup>9,10</sup> that intracellular buildup of VOCs has detrimental effects on membrane integrity. To test whether changes in VOC internal pools' distribution would lead to similar damaging effects despite an overall decrease in pool sizes in transgenic flowers, we performed staining with propidium iodide, which diffuses into cells and stains nucleic acids only if plasma membrane is disrupted<sup>38</sup>.
Indeed, nuclei were stained in some cells of 2-day-old *PhABCG12-9* petunia petals but not in those of wild-type (Supplementary Figure 10a). The similarity in round cell shape in transgenics and wild-type control, which does not have misshapen cells (Supplementary Figure 3), suggests VOC toxicity occurs in the conical cells.

Internal pools of VOCs with relatively higher vapor pressures such as benzaldehyde and methylbenzoate were more abundant in the wild-type cuticle relative to the rest of the cell (>90% of total internal pools, despite their small overall internal pool sizes) when compared to that of less volatile compounds, and this trend was conserved in transgenic lines (Extended Data Fig. 8a). Cuticular VOC concentration is determined by the equilibria at both the cuticle-atmosphere and cell wall-cuticle interfaces, which depend on compound relative volatility and octanol-water partition coefficient (as a proxy for wax-water partitioning), respectively. Comparisons of individual VOC distributions within the cell with corresponding compound volatility and octanol-water partition coefficient (Extended Data Fig. 8b and c) reveal that VOC volatility correlates with their partitioning in the cuticle. This suggests that the cuticle-atmosphere interface influences VOC distribution within the cell, consistent with the “pulling” effect of the atmosphere on compounds with relatively high volatility.

**Cuticle thickness affects rhythmicity of VOC emission.** VOC emission from petunia flowers occurs in rhythmic manner with highest flux at night$^{25,39}$. To dissect the effect of reduced cuticle thickness on VOC dynamics around peak of emission, floral volatiles and their internal pools were analysed in 2-day-old wild-type and *PhABCG12-9* flowers over 15 h period. In wild-type flowers, a rapid accumulation of internal pools was observed between 4 PM and 10 PM and remained constant until 4 AM (Extended Data Fig. 9a). This increase in VOC internal pools correlated with a notable peak in emissions (Extended Data Fig. 9b) between 10 PM and 1 AM,
and such profile was conserved for all individual VOCs (Supplementary Figure 11). In contrast, rhythmicity in VOC emission was practically absent in *PhABCG12-9* flowers (Extended Data Fig. 9b), while internal pools exhibited similar to wild type profile, but with ~27% lower maximum value (Extended Data Fig. 9a). Although transgenic petals still exhibited a clear temporal peak in VOC internal pools, the lack of a corresponding peak in emission suggests VOC redistribution upstream of the cuticle, thus dampening their release from the cell. Indeed, across six time points over a 15-h period on average ~ 23% less VOCs were present in cuticle of *PhABCG12-9* flowers than wild-type (Extended Data Fig. 9c), suggesting that limited cuticle sink for VOCs in transgenic flowers does not allow their rhythmic release.

**Effect of solvent-based dewaxing on rate of VOC emission.** In addition to the genetic approach, solvent-based dewaxing of petunia petals was used to examine the impact of reduced wax coverage on VOC emission without the potential influence of morphological changes observed in transgenic flower petals. These two approaches are complementary with their own strengths and limitations. Chemical dewaxing provides a snapshot of cuticle removal on VOC emission versus long-term changes in cuticle thickness in transgenics. Moreover, these two approaches have different starting points: in dewaxed flowers internal pools were instantaneously depleted, while in transgenic petals there is a constitutive shift in mass transfer resistance.

Dewaxing of wild type petals removed 70 ± 6% of wax leading to susceptibility to TB staining (Fig. 4a and b) and a decrease in total VOC emission and internal pools by over 80% at the end of the 4-h collection period (Fig. 4c). To ensure that the decrease in VOC production is not due to a loss of petal metabolic potential during dewaxing process, the activity of
phenylalanine ammonia lyase, which catalyzes the first committed step in the phenylpropanoid pathway, was measured before and 4 h after dewaxing and was found to be unchanged (Extended Data Fig. 10a). Moreover, feeding of dewaxed flowers with 150 mM Phe was able to recover both the VOC emission and biosynthetic fluxes observed in fed non-dewaxed flowers (Extended Data Fig. 10b and c). The fact that Phe feeding of dewaxed flowers was able to recover only 62 ± 10% of VOC internal pools (Extended Data Fig. 10d), despite a similar to non-dewaxed flowers absolute increase in internal pools upon feeding (1742 ± 52 nmol·g FW⁻¹ versus 1510 ± 277 nmol·g FW⁻¹ in non-dewaxed and dewaxed petals, respectively; Extended Data Fig. 10d and e), reflects the loss of the cuticular reservoir of VOCs. Taken together these results demonstrate that (i) flowers remain metabolically active 4 h after dewaxing and (ii) similar to PhABCG12-RNAi transgenic flowers removal of cuticle leads to decreased VOC production via feedback inhibition of Phe formation by internally accumulated volatile compounds. Indeed, carbon flux towards Phe was reduced after dewaxing. Although glucose was labeled 48% higher at 10 PM in dewaxed relative to non-dewaxed wild-type petals, the ¹³C labeling and pool sizes of Phe were both reduced by 84% and 38%, respectively, relative to non-dewaxed control (Supplementary Figure 8). In addition, the expression of Phe biosynthetic genes, ODO1 transcription factor, and PhABCG1 transporter was reduced upon dewaxing (Supplementary Figure 12) similar to the situation in PhABCG12-RNAi flowers (Supplementary Figure 9). This effect could be replicated by inducing internal VOC accumulation to toxic levels via feeding of wild-type flowers with 15 mM benzaldehyde¹⁰ (Supplementary Figure 13). Moreover, as in PhABCG12-RNAi flowers, dewaxed petals display plasma membrane disruption by internally accumulated VOCs, which is visible upon propidium iodine staining (Supplementary Figure 10b).
**Reduced cuticle thickness shifts mass transfer resistance.** Dewaxing of wild-type petunia petals led to an increase in VEF (Fig. 5a), as these flowers continue to emit VOCs, albeit at a low level, without changes in internal pools over the sampling time (Fig. 4c) further supporting the hypothesis that reduction of cuticle thickness decreases overall resistance. As petal dewaxing reduces VOC formation, to determine whether the high VEF value would be sustained under elevated VOC biosynthetic flux, dewaxed wild-type petunia petals were fed with 150 mM Phe. Unexpectedly, an increase in VOC biosynthesis decreased VEF (Fig. 5a), which could occur only if VOCs accumulate inside the cell in the absence of cuticle sink, thus demonstrating redistribution in the source of mass transfer resistance. VEF was also reduced in fed non-dewaxed wild-type petunia petals relative to the non-fed control and was very similar to the VEF of dewaxed wild-type petals fed with Phe (Fig. 5a). Comparisons of individual VOC distributions within the cuticle with and without Phe feeding revealed that compounds with low relative volatility accumulate internally under high biosynthetic flux (Fig. 5b), suggesting a shift in mass transfer resistance source upon increased VOC biosynthesis (Fig. 6).

To test if this effect was conserved in transgenic petals, *PhABCG12-9* RNAi flowers were subjected to dewaxing (Fig. 4a). Similar to wild-type flowers, dewaxing *PhABCG12-9* petals reduced emission and internal pool accumulation (Fig. 4d). Furthermore, dewaxing of transgenic petals increased VEF (Fig. 5a), but the value remained below 1 and was still lower than in control dewaxed petals. Thus, despite an 85% reduction in wax amount in *PhABCG12-9* petals upon dewaxing relative to non-dewaxed wild-type petals (Fig. 4a), VOCs still continue to accumulate internally (Fig. 4d). The inability to discharge the VOC internal pools upon combined RNAi reduction of cuticle thickness and dewaxing suggests that the resistance to mass transfer in transgenic flowers has shifted from the cuticle to other sources, such as the plasma
membrane and cell wall. Consistent with this conclusion, feeding of PhABCG12-9 flowers with 150 mM Phe did not affect VEF (Fig. 5a), and similar to wild-type petunia flowers, led to the redistribution of VOC internal pools of only compounds with low relative volatility (Fig. 5c).

Taken together, these results show that (i) cuticle serves as a significant resistance barrier for VOC emission; (ii) without enough cuticle present to act as a sink, VOCs accumulate internally, leading to inhibition of precursor formation to prevent toxicity; (iii) compounds with lower relative volatility experience greater resistance at the cuticle; and (iv) other barriers, due to a shift in the mass transfer resistance, control emission under high VOC biosynthetic flux.

## Discussion

The cuticle is the final barrier VOCs cross before being released into the atmosphere. Our results show that cuticle of petunia flowers holds > 50% of internal VOCs (Fig. 1a), thus acting as the main site of VOC storage, and serving as the major source of mass transfer resistance. However, it mainly provides resistance to VOCs with low relative volatility with negligible effect on highly volatile compounds, physiological emissions of which largely depend on their biosynthesis (Figs. 3 and 5; Extended Data Fig. 8). The latter compounds have small internal pools relative to their emission flux, and these pools are predominantly allocated to the cuticle (Extended Data Fig. 8a). To sustain efficient emission, VOCs have to accumulate in the cuticle and decreasing the ability of cuticle to act as a concentrator/sink not only reduces overall emission (Figs. 2d and 4c), but also eliminates rhythmicity in VOC release (Extended Data Fig. 9b and c and Supplementary Figure 11). The inability for VOCs, particularly those with low relative volatility, to build up in the cuticle due to a reduction in its thickness via RNAi downregulation of PhABCG12 transporter (Fig. 2b and c) as well as dewaxing (Fig. 4a) leads to a redistribution of internal VOC
pools (Figs. 3f and 5b; Extended Data Fig. 8a), thus resulting in cellular damage (Supplementary Figure 10). To mitigate this detrimental effect, cells reduce production of VOCs by decreasing biosynthesis of the precursor Phe (Fig. 6a and Supplementary Figures 8, 9, and 13), a feedback effect that was also observed in flowers after chemical dewaxing (Fig. 4 and Supplementary Figures 8 and 12) but could be bypassed by Phe feeding (Extended Data Fig. 10b and d). Finally, the decrease in cuticle thickness leads to a shift in mass transfer resistance to other internal barriers (i.e. plasma membrane and cell wall) (Fig. 6b), and this shift is more pronounced under high biosynthetic flux, likely reflecting the limited capacity of the plasma-membrane localized VOC transporter, PhABCG1¹⁰. Taken together, these results reveal that the cuticle, despite being a passive transport barrier, is nonetheless an integral member of the overall scent network.

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## Figure legends
**Fig. 1** | VOC distribution within cell and effect of *PhABCG12* downregulation on total wax amount in petunia flowers. **a**, Cellular VOC distribution in 2-day-old wild-type petunia flowers. Data are means ± S.E. (n = 4 biological replicates). **b**, *PhABCG12* mRNA levels determined by qRT-PCR in buds (3-4 cm) of wild type (WT), empty vector (EV) control and seven independent *PhABCG12*-RNAi lines. Data are shown as a percentage of *PhABCG12* expression in WT set as 100% and are presented as means ± S.E. (n = 3 biological replicates). **c**, Total wax levels in 2-day-old flowers of WT, EV control and the three transgenic lines with greatest reduction in *PhABCG12* expression (lines 7, 8 and 9). Data are means ± S.E. (n = 5 biological replicates). *P* values were determined two-way ANOVA with the Tukey's multiple comparisons test relative to the WT (black) and EV (blue) controls.

**Fig. 2** | Effect of *PhABCG12* downregulation on flower phenotype, cuticle properties and scent network in petunia flowers. **a**, Representative 2-day-old petunia flowers from WT, EV control and three independent *PhABCG12*-RNAi lines (7, 8 and 9) before (upper panel) and 4 h after staining with toluidine blue (lower panel). Scale bar, 1 cm. **b**, Representative transmission electron microscopy (TEM) pictures of cross-sections of epidermal conical cells from adaxial petal surface of 2-day-old WT and *PhABCG12-9* flowers. Scale bar, 200 nm. **c**, Cuticle thickness measured in conical epidermal cells of 2-day-old WT and *PhABCG12-9* flowers as shown in **b**. The box represents IQR of the data, whiskers show the minimum and maximum, and the line indicates the median (n = 675 measurements for each). *P* value was determined by unpaired two-tailed Student's *t*-test relative to control. Total emissions (**d**), total internal pools (**e**) and biosynthetic fluxes (**f**) in 2-day-old flowers from WT, EV control and three *PhABCG12*-RNAi lines 7, 8 and 9. Data are means ± S.E. (n = 6 biological replicates in **d**; in **e** n = 16, 10, 9, 6 and
11 biological replicates for WT, EV, lines 7, 8 and 9, respectively; and n = 4 biological replicates in **f**). *P* values in **d**, **e** and **f** were determined by two-way ANOVA with the Tukey's multiple comparisons test relative to the WT (black) and EV (blue) controls.

**Fig. 3** | **Effect of *PhABCG12* down-regulation on VEFs, emission and biosynthetic fluxes of representative VOCs and cellular distribution of VOCs in 2-day-old petunia flowers.** a, Overall VEF (the ratio of emission flux to the total biosynthetic flux) for WT, EV control and three independent *PhABCG12*-RNAi lines 7, 8 and 9. *P* values were determined by two-way ANOVA with the Tukey's multiple comparisons test relative to the corresponding WT (black) and EV (blue) controls. **b**, VEF of individual volatiles emitted by WT and *PhABCG12-9* flowers, shown in order of increasing compound volatility. *P* values were determined by unpaired two-tailed Student's *t*-test relative to the corresponding WT control. **c**, Emission and biosynthetic fluxes for benzaldehyde (BAld), methylbenzoate (MB) and (**d**) benzylbenzoate (BB) in WT (upper panels) and *PhABCG12-9* line (lower panels). **e**, Cellular VOC distributions in petunia flowers of EV control and three independent *PhABCG12*-RNAi lines 7, 8 and 9. *P* values were determined by unpaired two-tailed Student's *t*-test relative to the corresponding EV control.
All data are means ± S.E. (n = 4 biological replicates in **a**, **b**, and **e**; and n = 3 biological replicates in **c** and **d**).

**Fig. 4** | **Effect of dewaxing on corolla wax levels, cuticle permeability, VOC internal pools and emissions in 2-day-old petunia flowers.** **a**, Total wax levels before and after dewaxing of wild-type and *PhABCG12*-RNAi line 9 flowers. Data are means ± S.E (n = 4 biological
replicates). *P* values were determined by two-way ANOVA with the Tukey's multiple comparisons test. **b**, Representative non-dewaxed and dewaxed wild-type flowers 2 h after staining with toluidine blue. Scale bar, 1 cm. **c**, Emission fluxes and VOC internal pools over a 4-h period in non-dewaxed and dewaxed wild-type and *PhABCG12*-RNAi line 9 (**d**) flowers. Data are means ± S.E (n = 4 biological replicates).

**Fig. 5** | Effect of dewaxing and Phe feeding on VEFs and cellular VOC distributions in 2-day-old petunia flowers. **a**, Overall VEFs for control (non-dewaxed) and dewaxed wild-type and *PhABCG12*-RNAi line 9 flowers with and without feeding with 150 mM Phe over 4 h period from 6 PM till 10 PM. Data are means ± S.E. (n = 4 biological replicates). *P* values were determined by two-way ANOVA with the Tukey's multiple comparisons test. **b**, Cellular distributions of total (left panel) and individual VOCs (right panel) in control (non-dewaxed) wild type and (c) *PhABCG12-9* line flowers with and without Phe feeding. Data are means ± S.E. (n = 4 biological replicates). *P* values were determined by unpaired two-tailed Student's *t*-test relative to the corresponding unfed control.

**Fig. 6** | Schematic presentation of shift in the mass transfer resistance sources upon reduction in cuticle thickness. **a**, Schematic representation of VOC (red icons) feedback effect on transcription of Phe biosynthetic genes. Black arrows represent enzymes, orange arrows represent RNA translation, and green arrows represent gene transcription. Red lines show feedback inhibition of Phe biosynthetic gene transcription by accumulating VOCs via unknown mechanisms. **b**, Overall resistance shown by a resistance-in-series model. The barrier with the
highest contribution to overall resistance to VOC (red icons) mass transfer out of the cells is shown by the largest resistor symbol. Purple icon in plasma membrane represents previously characterized PhABCG1 transporter¹⁰.

# Methods

## Plant materials and growth conditions

Wild-type and transgenic *Petunia hybrida* cv. Mitchell diploid (W115; Ball Seed Co., West Chicago, IL) were grown under standard greenhouse conditions with a light period from 6:00 h to 21:00 h. Both *PhABCG11*-RNAi and *PhABCG12*-RNAi constructs were synthesized by Genscript (Piscataway, NJ). The *PhABCG11*-RNAi construct contained two spliced *PhABCG11* cDNA fragments corresponding to nucleotides 1234-1733 (in sense orientation) and 1234-1533 (in antisense orientation to form a hairpin structure), while *PhABCG12*-RNAi construct included two spliced *PhABCG12* cDNA fragments corresponding to nucleotides 225-724 (in sense orientation) and 225-524 (in antisense orientation), and both constructs were synthesized with flanking *AttL1* and *AttL2* sequences. Before synthesis, both the *PhABCG11* (*PhABCG11a* in Supplementary Table 1) and *PhABCG12* (*PhABCG12b* in Supplementary Table 1) constructs were verified to target only the desired genes and not to lead to off-target interference using the Sol Genomics Network VIGS Tool (http://vigs.solgenomics.net/). The analysis was performed by comparing the *PhABCG11a* and *PhABCG12b* RNAi target sequences against the *P. axillaris* and *P. inflata* (parents of *P. hybrida*) genomes³⁴ using default parameters, except that the n-mer setting was adjusted to 24 nucleotides, which is generally considered to be below the threshold to efficiently trigger down regulation⁴⁰,⁴¹. This analysis revealed that no other genes would be targeted by the *PhABCG11* and *PhABCG12* dsRNA triggers. In addition, the maximum stretch
of uninterrupted shared sequence bewteen *PhABCG12* and *PhABCG11* as well as *PhABCG1* was found to be 12 nucleotides for each gene (Supplementary Figure 2a). Further analysis of *PhABCG12b* RNAi down-regulation on expression of *PhABCG12* homologs in *PhABCG12*-RNAi lines determined by qRT-PCR with gene-specific primers revealed that their expression remained unchanged (Supplementary Figure 2b). Each of the synthetic RNAi fragments was placed under the control of the petal development-specific MYB1 promoter from Japanese morning glory (*Ipomoea nil*) (pDONRG-P4PIR-InMYB1pro vector the kind gift of Dr. Y. Oshima)³⁶ in the binary destination vector R4pGWB5_stop_HSP⁴² using the MultiSite Gateway Three-Fragment Vector Construction Kit (Invitrogen, Carlsbad, CA). Transgenic *P. hybrida* plants were obtained via *Argobacterium tumefaciens* (strain GV3101 carrying the final *PhABCG11*-RNAi or *PhABCG12*-RNAi constructs) leaf disc transformation using a standard transformation protocol⁴³.

## Cuticular wax extraction and analysis

Waxes were extracted from petals of 2-day-old wild-type and transgenic petunia flowers at 6 PM by submerging corollas in 10 mL of HPLC-grade hexane for 30 sec. The solvent was decanted into a fresh glass vial and dried completely under a gentle stream of N₂. The dried wax residues were derivatized using 200 µL of *N,O*-bis(trimethylsilyl)trifluoroacetamide (BSTFA Derivatization Grade, Sigma-Aldrich, St. Louis, MO) and pyridine (1:1 BSTFA:pyridine, v/v) for 15 min at 100°C, and excess BSTFA and pyridine was evaporated under N₂. Derivatized waxes were solubilized in hexane containing a known quantity of the internal standard *n*-tetracosane (99%, Sigma-Aldrich, St. Louis, MO) and samples were analyzed on an Agilent 6890 gas chromatograph (GC) equipped with a HP5-MS column (0.25mm x 30m x 0.25µm,
Agilent, Santa Clara, CA) and coupled to a 5975B inert MSD quadrupole mass spectrometer.
The GC separation conditions were: an initial 2 min hold at 40°C, followed by a 15°C/min
temperature ramp to 200°C and held for 2 min, then increased at 3°C/min to 320°C, and held for
15 min. Compounds were identified based on retention times and fragmentation patterns, and
quantified by peak areas relative to the internal standard. Specific correction (response) factors
and calibration curves were developed from external standards of fatty acids, primary alcohols,
and *n*-alkanes of varying chain length and composition. Total wax quantity was obtained by
pooling the wax extracts from 20 to 30 flowers in independent replicates, and measuring the
weight of the dried residues.

RNA extraction and qRT-PCR analysis

Sample collection, RNA extraction, and qRT-PCR was carried out as described previously⁴⁴ with
minor modifications. Briefly, RNA was isolated from petals collected at the indicated
developmental stages and times of the day using the Spectrum Plant Total RNA Kit (Sigma-
Aldrich, St. Louis, MO, USA). Total RNA (2 µg) was treated by DNase I (Promega, Madison,
WI, USA) before reverse transcription to cDNA using a 5x All-In-One RT MasterMix (Applied
Biological Materials, Richmond, BC, Canada). Changes in *PhABCG11* and *PhABCG12*
expression during flower development were analyzed by qRT-PCR using elongation factor 1-α
(EF1-α) as a reference gene⁴⁵,⁴⁶. Relative quantification of *PhABCG1*, *PhABCG11*, *PhABCG12*,
*PhDAHPS*, *PhEPSPS*, *PhCM1*, *PhCM2*, and *PhODO1* transcripts were performed by qRT-PCR
relative to the reference gene *PhUBQ10* ¹⁰,⁴⁶. qRT-PCR data were collected using StepOne
v.2.2.2. and analyzed according to the 2⁻⁰⁻²ΔΔCt method⁴⁷. The absolute quantification of gene
transcript levels was performed based on standard curves generated by qRT-PCR with gene-
specific primers using respective purified cDNA fragments diluted to several concentrations
between 1.3 pg ml⁻¹ to 24.3 ng ml⁻¹ for *PhABCG11* and 32.4 pg ml⁻¹ to 608.0 ng ml⁻¹ for *PhABCG12*. Absolute quantities of individual transcripts were calculated and expressed as pictograms (pg) of mRNA per 200 ng of total RNA. Primers used for qRT-PCR are listed in Supplementary Table 4.

### Transmission electron microscopy

Small pieces (1 mm in diameter) of wild type and transgenic petals on day 2 postanthesis were fixed in 2% paraformaldehyde and 2% glutaraldehyde in 0.1 M sodium cacodylate buffer. After primary fixation, samples were rinsed and post-fixed in buffered 1% osmium tetroxide containing 0.8% potassium ferricyanide. Subsequently, samples were dehydrated with a graded series of ethanol, transferred into acetonitrile and embedded in EMbed-812 resin. Thin sections (80 nm) were prepared on a Leica UC6 ultramicrotome and stained with 4% uranyl acetate and lead citrate. Sections were imaged on a FEI Tecnai T12 electron microscope equipped with a tungsten source and operating at 80 kV. Images were acquired in DM4 format on a Gatan Orius SC200 camera and converted to obtain TIFF files. To analyze cuticle thickness in conical epidermal cells from transgenic and WT petals, cuticle thickness shown in TEM images was measured by ImageJ software version 1.52a⁴⁸.

### Cryo-scanning electron microscopy (cryo-SEM)

Wild-type and transgenic petals on day 2 postanthesis were mounted with OCT cryo-sectioning media on a flat specimen holder and cryo fixed by immersion in a nitrogen slush using a Gatan Alto 2500 Cryo-Preparation System (Gatan Inc. Pleasanton, CA). Samples were immersed until liquid nitrogen boiling ceased, and then cryo transferred into the Gatan preparation chamber set
at -170°C. Samples were then immediately inserted into the SEM cryo stage set at -90°C to
sublimate surface frost and for initial viewing. Once frost free, samples were returned to the
cryo-preparation chamber, allowed to cool and halt sublimation before sputter coating with a
platinum target with 18 mA for 120 sec at -170°C. Final imaging was done at -140°C with an
accelerating voltage of 5 kV and spot size 3 on the Nova Nano 200 SEM (Hillsboro, OR).

Collection and analysis of plant volatiles

Floral volatile emission was analyzed by dynamic headspace collection of volatiles at 25°C on
columns containing 50 mg Poropak Q (80–100 mesh) (Sigma-Aldrich) and analyzed by GC-MS
as described previously¹⁰,⁴⁹. Briefly, emitted floral volatiles were collected from detached
transgenic and control flowers on day 2 postanthesis from 6 PM to 10 PM. Subsequently, ~280
µL dichloromethane containing 52 nmol of internal standard (naphthalene) was used to elute
volatiles from collection columns for GC-MS analysis. Internal pools of volatiles from
transgenic and control tissues collected at 22:00 h on day 2 postanthesis were extracted
according to ¹⁰ and quantified by GC-MS as described previously⁴⁹. Briefly, 0.5 g of petal tissue
was collected, ground to a fine powder in liquid nitrogen, and internal pools were extracted in 5
mL of dichloromethane containing 52 nmol of internal standard (naphthalene) for 12 h at 4°C
with shaking. Samples were concentrated to ~ 200 µL under a stream of nitrogen gas before
analysis by GC-MS. All GC-MS data were collected and analyzed using Agilent MassHunter
Workstation Software v.B. 07.01.

Toluidine blue staining
Toluidine blue staining was performed following published protocol as follows³⁷,⁵⁰. Excised petals of petunia flowers on day 2 postanthesis collected at 3 PM were submerged in 0.05% toluidine blue aqueous solution. Flowers were incubated for 2 or 4 h, as indicated, before imaging.

## Water loss measurement

Water loss measurements were carried out according to ⁵¹,⁵² with some modifications. 2-day-old petunia flowers (3 flowers per replicate) were detached and left in ambient laboratory conditions. The fresh weight of the flowers was recorded at the indicated times over 24 h to obtain total water loss rate.

## Starch and sucrose quantification

Starch was extracted as described previously⁵³. In brief, corolla tissue (collected on day 2 postanthesis at 3 PM) was flash frozen in liquid nitrogen, ground to fine powder, and 100 mg of powder was suspended in 1 ml 80% ethanol by vortexing. After heating at 100°C for 3 min, samples were centrifuged at 4000 g for 15 min at room temperature, and the supernatant was discarded. This process was repeated five additional times. The final starch-containing pellet was resuspended in 1 ml water and the starch gelatinized by heating for 10 min at 100°C. Starch was hydrolyzed by adding sodium acetate (pH 5.5) to a final concentration of 100 mM, along with 6 units of α-amyloglucosidase and 0.5 units of α-amylase, and incubating at 37°C for 4 h. The hydrolyzed starch samples were centrifuged at 10,000 g for 5 min at room temperature, and a 20-µl sample was subjected to chromatographic separation using an Agilent 1260 Infinity II HPLC
system and a Hi-Plex Ca column (300 × 7.7 mm; Agilent) held at 80°C with water as the isocratic mobile phase. Glucose was detected using an Agilent 1260 Refractive Index Detector.

Sucrose was extracted and analyzed as described previously⁵⁴. In brief, corolla tissue (collected on day 2 postanthesis at 3 PM) was flash frozen in liquid nitrogen and ground to fine powder. 0.5 ml of methanol containing 1 mg/ml of sorbitol as internal standard was used to extract 100 mg of the homogenized tissue at 70°C for 15 min. After cooling, 0.5 ml of water was added and the sample was incubated at room temperature for an additional 15 minutes with periodic vortexing. Samples were centrifuged at 12,000 g for 15 min at room temperature, and 20 µl of the supernatant was dried under a gentle flow of nitrogen. The resulting residue was derivatized by adding 50 µl of pyridine containing 20 mg/ml methoxyamine hydrochloride and incubating for 30 min at 30°C, followed by additional 90 min incubation at 37°C in the presence of 50 µl MSTFA. Derivatized samples were analyzed by GC-MS by introducing 1 µl of sample into the GC injector held at 230°C in split mode using a split ratio of 1:4. The column temperature was held at 150°C for 1 min and then increased at 5°C min⁻¹ to 175°C, 1°C min⁻¹ to 185°C, and 10°C min⁻¹ to 250°C.

## Respiration rate measurements

Petal respiration rates were measured using an infrared gas analyser (LI-6800, Licor, Lincoln, NE, USA). Petals of flowers on plants grown in greenhouse conditions were enclosed in the cuvette and CO₂ efflux was measured after stability of environmental parameters in the cuvette. Conditions in the cuvette were maintained as close to glasshouse environmental conditions as possible, a light intensity was set a 1000 µmol quanta m⁻² s⁻¹, temperature was maintained at 25°C and vapour pressure difference controlled at 1.2 kPa. Respiration rates were measured for
each petal after gas exchange parameters stabilized, which occurred within approximately 5 min
after enclosure of petal inside the cuvette. All data were standardized against petal area in the
cuvette.

## Seed germination assays

Seeds of petunia wild-type and three independent *PhABCG12*-RNAi lines (7, 8 and 9) (210 seeds per line) were sterilized in 75% ethanol, rinsed in sterile water for five times and placed on plates containing MS agar with 3% sucrose (42 seeds per plate). After cold stratification at 4°C for 5 d, seed were transferred to a growth room (21°C, 8-h dark/22°C 16-h light). The radicle emergence was considered as germination. The number of germinated seeds was recorded every 24 h within 7 days.

## Analysis of seed yield

Seed yield was measured as described previously⁵⁵ with minor modifications. Briefly, flowers were cross-pollinated at 6 PM on day 2 postanthesis. After 4-5 weeks, mature seed pods with brown color were collected individually and placed in a Office Depot® Brand Coin Envelope (5.7 cm × 8.9 cm). Seeds were collected and dried for 2 d at 37°C after pods were cracked. Subsequently, the weight of total seeds per pod, seed weight and total number of seeds per pod were measured.

## Analysis of chlorophyll levels

Chlorophyll levels were measured according to ⁵⁶ with minor modifications. Briefly, wild-type and transgenic petals collected at 3 PM on day 2 postanthesis were ground to fine powder in
liquid nitrogen and chlorophylls were extracted in dim light by adding twice 500 µl of 80% aqueous acetone to 100 mg of sample powder. After centrifugation at 12,000 g for 10 min, supernatants were combined and the concentration of chlorophyll a and b were determined by spectrophotometrically at 663 nm and 645 nm, respectively, using a Nanodrop 2000c spectrophotometer (Thermo Scientific, Waltham, MA). The total chlorophyll levels were calculated according to⁵⁶.

## Propidium iodide staining and confocal laser scanning microscopy

Petals of wild-type and *PhABCG12*-RNAi flowers collected at 3 PM on day 2 postanthesis were stained in propidium iodide solution (10 µg/mL, Invitrogen, Carlsbad, CA) for 1 h with shaking at room temperature. Confocal fluorescence microscopy was performed as described previously¹⁰,⁵⁷ with minor modifications. Briefly, propidium iodide-stained samples were excited with a 561-nm laser and images were recorded by a 617/73 emission filter using a Zeiss LSM 880 Upright Confocal system and operated using Zeiss Efficient Navigation (ZEN)-Black Edition software (Zen 2.3 SP1, Carl Zeiss Inc., Oberkocken, Germany).

## Dewaxing of petunia petals

To determine the quantity of volatiles in petunia epicuticular waxes, petals on day 2 postanthesis were rapidly dipped in hexane (< 5 seconds) and then were lightly dried under the fume hood to remove excess hexane. Care was taken to prevent full submersion of petals so that only the adaxial surface of the flower touched the solvent. Hexane pools were independently profiled for volatiles and wax quantity using GC-MS as described above, while dewaxed petals were
sampled for VOC internal pools and emissions as described above. Total cuticular volatile
fractions were determined according to the formula:

$$
\text{Cuticular VOC fraction} = \frac{C_i^{\text{hex}}}{C_i^{\text{total}}} \cdot \frac{1}{R} \qquad (\text{equation 1})
$$

where $C_i^{hex}$ is the volatile pool in the hexane fraction, $C_i^{total}$ is the total volatile pool, and $R$ is the fraction of the total wax recovered during the dewaxing.

To test whether dewaxed flowers retained biosynthetic capability, dewaxed wild-type
petals were fed with 15 and 150 mM Phe and VOC internal pools and emission were
subsequently analyzed between 6 PM and 10 PM. In addition, PAL activity was measured in
dewaxed and non-dewaxed wild-type petals on day 2 postanthesis as described previously³⁹,⁵⁴,
except that assays were adapted to high-performance liquid chromatography (HPLC) detection
of cinnamic acid formation from L-Phe. In brief, 100 µl reaction mixture containing protein
samples from dewaxed and non-dewaxed petals (5-30 µg), 2 mM L-Phe, and 0.1 M sodium
borate (pH 8.8) was incubated at room temperature for 1 h and then terminated by adding 5 µl of
6 N HCl. After centrifugation, 10 µl of the supernatant was analyzed by HPLC equipped with the
Agilent Poroshell 120 EC-C18 column (3.0 × 150 mm × 2.7 µm) held at 35°C using an 8-min
linear gradient of 5-70% acetonitrile in 0.1% formic acid at a flow rate of 0.4 mL min⁻¹.
Cinnamic acid was detected by UV absorbance at 270 nm and quantified using Open LAB CDS
Chemstation Rev. C.01.08[210] and authentic standard. Control assays including (i) with all
reaction components with boiled protein extracts; (ii) with all reaction components except
protein extracts, and (iii) with all components except for L-Phe, did not produce cinnamic acid.

Metabolic flux analysis with ¹³C₆-phenylalanine labeling
Wild-type and PhABCG12-RNAi-9 petunia corollas excised from flowers 2 days postanthesis were placed in a cut 1.7-ml eppendorf tube contain 300 µl solution of 150 mM ring-labeled ¹³C₆-Phe (Cambridge Isotope Laboratories, Andover, MA). Emitted and internal pools of volatiles were collected between 6 PM -10 PM as described above at 30-, 60-, 120-, and 240-min time points after beginning of feeding and analyzed by GC-MS. Labeled volatiles were identified by M+6 and M+12 (for VOCs with two aromatic rings) shifts in *m/z* and labeling percentage was determined based on the peak areas of labeled and unlabeled volatiles. Metabolic fluxes were calculated by applying mass balances and *in vivo* metabolite labeling profiles as described previously²⁴. Emission fluxes and time-dependent accumulation of internal pools were estimated using linear regression, with the variances of the estimated slopes calculated using the standard linear regression procedure described previously⁵⁸. As fluxes are a function of the linear regression and other experimental measurements, flux variances are derived by considering the propagation of errors based on the following equation:

$$ \sigma_y^2 = \sum_{i=1}^{n} \sigma_{x_i}^2 \left( \frac{\partial y}{\partial x_i} \right)^2 \qquad (\text{equation 2}) $$

where $y = f(x₁, ..., xₙ)$.

## Benzaldehyde feeding

Wild-type petunia corollas on day 2 postanthesis were collected at 9 AM and fed with 0 and 15 mM of benzaldehyde for 6 h, as described previously¹⁰. Mock and benzaldehyde-fed petunia corollas were collected at 3 PM, ground to a fine powder in liquid nitrogen, and used for subsequent RNA extraction, cDNA synthesis, and qRT-PCR analysis as described above.

## Stable isotope labeling with ¹³C-glucose
Corollas from flowers 2 days postanthesis were placed, cut surface down, on filter paper moistened with 3% (w/v) uniformly labelled ¹³C₆-glucose (Cambridge Isotope Laboratories, Inc. Tewksbury, MA, USA) at 6 PM. After 1, 2, and 4 h glucose was extracted and analysed as described for sucrose above. Phe was extracted and analysed as described previously⁵⁹ with some modification as follows. Tissue was extracted with 10 µl/mg of 75% methanol (v/v) containing 1 µg/ml of *p*-fluoro-DL-phenylalanine as internal standard. After a 30-min incubation at 65°C, samples were centrifuged at 15,000 g for 15 min. The supernatant was transferred to a separate tube, dried to completeness in a Labconco CentriVap concentrator (Kansas City, MO, USA), resuspended in ultrapure water with 20% (v/v) methanol and 10 µL of sample was injected into LC-MS/MS. The incorporation of the ¹³C labelled glucose in the shikimate pathway was estimated by determining the % ¹³C label in phenylalanine via reverse-phase chromatography on a Shimadzu HPLC-20 AD (Shimadzu, Columbia, MD, USA) coupled to a AB Sciex 5500 mass spectrometer (AB Sciex, Redwood City, CA, USA) with electrospray ionization in negative ion mode. The ionization parameters were manually tuned and are listed in the Supplementary Table S5. Chromatographic separations were performed on a 150 mm x 4.6 mm Zorbax Eclipse C8 column, with 5µm particles (Agilent Technologies, Santa Clara, CA), at a column temperature 30°C and a flow rate of 1 ml/min. A linear gradient of aqueous solvent A (2.5 mM ammonium acetate in ultrapure water, adjusted to pH 5.3 with acetic acid) and organic solvent B (97.8% acetonitrile, 2% ultrapure water, and 0.2% formic acid) was as follows: 10% B (v/v) for 1 min, 10-20% B over 6 min, 20-20.8% B over 9 min, 20.8-50% B over 1 min, 50-70% B over 1 min, hold at 70% B for 3 min, return to 10% B over 2 min followed by equilibration at 10% B for 3 mins for a total of 26 min. The peaks were integrated and quantified by Analyst 1.5.1 software.
For quantifying %C labelling, all the possible Q1/Q3 combinations were considered. Labelling of both glucose and Phe were corrected for natural isotope abundance.

## Calculation of volatile emission factor

Biosynthetic and emission fluxes were estimated from dynamic sampling of volatile internal pools and emissions from detached transgenic and control petunia flowers on day 2 postanthesis from 6 PM to 10 PM. Time-dependent accumulation of volatile internal pools and the rate of emission were estimated using linear regression, with propagation of regression and experimental errors calculated using equation 2. Biosynthetic flux was determined from the mass balance for an individual volatile:

$$ \frac{dC_i}{dt} = v_{syn,i} - v_{emi,i} \quad (equation 3) $$

where $v_{syn,i}$ and $v_{emi,i}$ are the biosynthetic and emission fluxes of a volatile compound i, respectively. Emission and biosynthetic fluxes for individual compounds are shown in Supplementary Figure 14. The volatile emission factor (VEF) is the ratio between the emission and biosynthesis of a volatile compound:

$$ VEF_i = \frac{v_{emi,i}}{v_{syn,i}} \quad (equation 4) $$

The overall VEF is determined as the ratio between the sum of all individual volatile emission fluxes and the sum of all individual volatile biosynthetic fluxes:

$$ VEF = \frac{\sum_{i=1}^{n} v_{emi,i}}{\sum_{i=1}^{n} v_{syn,i}} \quad (equation 5) $$

where n denotes the number of volatiles detected in the pools and headspace of petunia flowers.
**Measurement of flower fresh weight, corolla diameter and corolla fresh weight**

Flower fresh weight and corolla diameter of wild type, empty vector control and 3 independent *PhABCG12*-RNAi lines (7, 8 and 9) were measured on 2-day-old detached flowers collected at 6 PM. Corolla fresh weight of all the above lines was recorded at 10 PM for 2-day-old detached flowers after scent collection.

**High humidity experiment**

To test whether the phenotypic changes in *PhABCG12*-RNAi lines were caused by water stress, 2-month-old wild-type and 3 independent *PhABCG12*-RNAi lines (7, 8 and 9) were moved from standard greenhouse conditions with a normal humidity (< 50%) to a growth chamber with a high relative humidity (75%) and allowed to acclimate for 1 week. Then, 2-day-old flowers were collected at 6 PM and flower fresh weight was recorded as well as total VOC emission and internal pools were analyzed as described before and compared to that of control 2-day-old flowers growing in the standard greenhouse conditions with normal humidity.

**Data Availability Statement**

The authors declare that all the data supporting the fundings of this study are available within the paper and its supplementary information files or from the corresponding author upon reasonable request. Source data are provided with this paper. *Petunia axillaris* genomic data were obtained from http://solgenomics.net using *Petunia axillaris* v1.6.2 genome database.

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## Acknowledgements

This work was supported by grant from the National Science Foundation IOS-1655438 to N.D. and J.A.M. and by the USDA National Institute of Food and Agriculture Hatch Project number 177845 to ND. The authors acknowledge the use of the imaging facilities of the Bindley Bioscience Center, a core facility of the NIH-funded Indiana Clinical and Translational Sciences
Institute, for collection of confocal microscopy images. We thank Dr. Y. Oshima (National Institute of Advanced Industrial Science and Technology, Japan) for providing pDONOR_P4PIR-InMYB1pro and R4pGWB5_stop_HSP vectors. We thank R. Seiler and L. Mueller for technical assistance on SEM and TEM, respectively.

## Authors contributions

B.B., J.A.M., and N.D. conceived the project and designed research; P.L. performed analysis of flower phenotypes, protein levels, seed production and germination rates, VOC emission and their internal pools, internal cellular VOC distribution over 15 h period, toluidine and propidium iodide staining experiments, expression analysis, and feeding experiments. S.R. performed wax profiling, VOC distribution and metabolic flux analyses, dewaxed experiments, water loss determination, and analyzed rhythmicity of VOC production and accumulation. B.B. generated PhABCG12-RNAi lines and performed initial expression and metabolic profiling. J.H.L. performed analysis of sucrose and starch levels as well as flux through the shikimate pathway. A.D. run and analyzed samples for shikimate pathway flux experiments. S.M. analyzed respiration rate. P.L., S.R, B.B., J.H.L., A.D., S.M., J.A.M., and N.D. analyzed data; N.D. wrote the manuscript with contribution from all authors. All authors read and edited the manuscript.

## Competing interests

The authors declare no competing financial interests.
<figure><img src="image_2.png" /><figcaption>**Fig. 1** | VOC distribution within cell and effect of *PhABCG12* downregulation on total wax amount in petunia flowers. **a**, Cellular VOC distribution in 2-day-old wild-type petunia flowers. Data are means ± S.E. (n = 4 biological replicates). **b**, *PhABCG12* mRNA levels determined by qRT-PCR in buds (3-4 cm) of wild type (WT), empty vector (EV) control and seven independent *PhABCG12*-RNAi lines. Data are shown as a percentage of *PhABCG12* expression in WT set as 100% and are presented as means ± S.E. (n = 3 biological replicates). **c**, Total wax levels in 2-day-old flowers of WT, EV control and the three transgenic lines with greatest reduction in *PhABCG12* expression (lines 7, 8 and 9). Data are means ± S.E. (n = 5 biological replicates). *P* values were determined two-way ANOVA with the Tukey's multiple comparisons test relative to the WT (black) and EV (blue) controls.</figcaption></figure>
<figure><img src="image_3.png" /></figure>

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<figure><img src="image_9.png" /><figcaption>**Fig. 2** | Effect of *PhABCG12* downregulation on flower phenotype, cuticle properties and scent network in petunia flowers. **a**, Representative 2-day-old petunia flowers from WT, EV control and three independent *PhABCG12*-RNAi lines (7, 8 and 9) before (upper panel) and 4 h after staining with toluidine blue (lower panel). Scale bar, 1 cm. **b**, Representative transmission electron microscopy (TEM) pictures of cross-sections of epidermal conical cells from adaxial petal surface of 2-day-old WT and *PhABCG12-9* flowers. Scale bar, 200 nm. **c**, Cuticle thickness measured in conical epidermal cells of 2-day-old WT and *PhABCG12-9* flowers as shown in **b**. The box represents IQR of the data, whiskers show the minimum and maximum, and the line indicates the median (n = 675 measurements for each). P value was determined by unpaired two-tailed Student's t-test relative to control. Total emissions (**d**), total internal pools (**e**) and biosynthetic fluxes (**f**) in 2-day-old flowers from WT, EV control and three *PhABCG12*-RNAi lines 7, 8 and 9. Data are means ± S.E. (n = 6 biological replicates in **d**; in **e** n = 16, 10, 9, 6 and 11 biological replicates for WT, EV, lines 7, 8 and 9, respectively; and n = 4 biological replicates in **f**). P values in **d**, **e** and **f** were determined by two-way ANOVA with the Tukey's multiple comparisons test relative to the WT (black) and EV (blue) controls.</figcaption></figure>
<figure><img src="image_10.png" /><figcaption>**Fig. 3** | Effect of *PhABCG12* down-regulation on VEFs, emission and biosynthetic fluxes of representative VOCs and cellular distribution of VOCs in 2-day-old petunia flowers. a, Overall VEF (the ratio of emission flux to the total biosynthetic flux) for WT, EV control and three independent *PhABCG12*-RNAi lines 7, 8 and 9. *P* values were determined by two-way ANOVA with the Tukey's multiple comparisons test relative to the corresponding WT (black) and EV (blue) controls. b, VEF of individual volatiles emitted by WT and *PhABCG12*-9 flowers, shown in order of increasing compound volatility. *P* values were determined by unpaired two-tailed Student's *t*-test relative to the corresponding WT control. c, Emission and biosynthetic fluxes for benzaldehyde (BAld), methylbenzoate (MB) and (d) benzylbenzoate (BB) in WT (upper panels) and *PhABCG12*-9 line (lower panels). e, Cellular VOC distributions in petunia flowers of EV control and three independent *PhABCG12*-RNAi lines 7, 8 and 9. *P* values were determined by unpaired two-tailed Student's *t*-test relative to the corresponding EV control. All data are means ± S.E. (n = 4 biological replicates in a, b, and e; and n = 3 biological replicates in c and d).</figcaption></figure>
<figure><img src="image_11.png" /></figure>

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**Fig. 4** | Effect of dewaxing on corolla wax levels, cuticle permeability, VOC internal pools and emissions in 2-day-old petunia flowers. **a**, Total wax levels before and after dewaxing of wild-type and *PhABCG12*-RNAi line 9 flowers. Data are means ± S.E (n = 4 biological replicates). *P* values were determined by two-way ANOVA with the Tukey's multiple comparisons test. **b**, Representative non-dewaxed and dewaxed wild-type flowers 2 h after staining with toluidine blue. Scale bar, 1 cm. **c**, Emission fluxes and VOC internal pools over a 4-h period in non-dewaxed and dewaxed wild-type and *PhABCG12*-RNAi line 9 (**d**) flowers. Data are means ± S.E (n = 4 biological replicates).
<figure><img src="image_17.png" /></figure>

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**Fig. 5** | Effect of dewaxing and Phe feeding on VEFs and cellular VOC distributions in 2-day-old petunia flowers. **a**, Overall VEFs for control (non-dewaxed) and dewaxed wild-type and *PhABCG12*-RNAi line 9 flowers with and without feeding with 150 mM Phe over 4 h period from 6 PM till 10 PM. Data are means ± S.E. (n = 4 biological replicates). *P* values were determined by two-way ANOVA with the Tukey's multiple comparisons test. **b**, Cellular distributions of total (left panel) and individual VOCs (right panel) in control (non-dewaxed) wild type and (c) *PhABCG12-9* line flowers with and without Phe feeding. Data are means ± S.E. (n = 4 biological replicates). *P* values were determined by unpaired two-tailed Student's *t*-test relative to the corresponding unfed control.
a

<figure><img src="image_20.png" /></figure>

b

Wild type

PhABCG12-RNAi

<figure><img src="image_21.png" /></figure>

<figure><img src="image_22.png" /><figcaption>**Fig. 6** | Schematic presentation of shift in the mass transfer resistance sources upon reduction in cuticle thickness. **a**, Schematic representation of VOC (red icons) feedback effect on transcription of Phe biosynthetic genes. Black arrows represent enzymes, orange arrows represent RNA translation, and green arrows represent gene transcription. Red lines show feedback inhibition of Phe biosynthetic gene transcription by accumulating VOCs via unknown mechanisms. **b**, Overall resistance shown by a resistance-in-series model. The barrier with the highest contribution to overall resistance to VOC (red icons) mass transfer out of the cells is shown by the largest resistor symbol. Purple icon in plasma membrane represents previously characterized PhABCG1 transporter<sup>10</sup>.</figcaption></figure>