Milkweeds (Asclepias spp.) are a classic system for studying plant chemical defences. This plant family has evolved a number of physical and chemical defences against a suite of generalist and specialist herbivores (Agrawal & Fishbein 2006). Milkweeds are often defended by cardenolides, bitter-tasting compounds that elicit aversive or emetic responses in both vertebrates (Brower et al. 1968) and invertebrates (Dussourd & Hoyle 2000). Cardenolides affect animals by inhibiting the ubiquitous sodium–potassium cellular pump Na+/K-ATPase (Malcolm 1991). They are poisonous to generalists at very low doses, yet can be tolerated and even co-opted for use as a chemical defence by specialists (Dussourd & Hoyle 2000; Agrawal et al. 2012).
Cardenolide biosynthesis, in which sterol precursors are modified into the 5b cardenolide genins that form the backbone of each compound, has not been fully characterized in plants and is hypothesized to depend on more than one pathway (reviewed in Agrawal et al. 2012). Cardenolide activity is primarily determined by the steroid nucleus and its stereochemistry, but the lactone and sugar side chains can modify compound selectivity and interactions with Na+/K-ATPase (Repke 1985; Paula, Tabet & Ball 2005). Structural differences, such as the number of glucose, CH3 or OH groups, between individual compounds affect polarity, toxicity and rate of absorption by animals postconsumption (Malcolm 1991; Agrawal et al. 2012).
The concentrations of cardenolides in leaves vary substantially across the genus Asclepias (Agrawal & Fishbein 2006; Agrawal, Lajeunesse & Fishbein 2008) and can also vary among plant parts such as the leaves, roots, pith and epidermis within the same plant (Nelson, Seiber & Brower 1981; Fordyce & Malcolm 2000; Rasmann et al. 2009). Although Asclepias nectar is reportedly toxic to honeybees (Pryce-Jones 1942), cardenolides have not previously been reported in the floral nectar.
The genus Asclepias is a monophyletic group (Agrawal & Fishbein 2008) composed of about 135 species found in the Americas (Woodson 1954; Agrawal & Fishbein 2008; Fishbein et al. 2011). We selected 12 closely related species in the series Incarnatae (Fig. 1, Table 1), as this is a monophyletic group with relatively well-resolved phylogenetic relationships (Agrawal & Fishbein 2008) and significant variation in leaf cardenolide concentrations among species (Agrawal, Lajeunesse & Fishbein 2008). Each plant was grown from seed and maintained in one of two greenhouse collections at Cornell University; seeds were collected by the authors, colleagues or from native plant nurseries and, once in bloom, we used flowers to verify species identity.
Figure 1. Average gross cardenolide concentrations (±SE) for (a) nectar, and (b) leaves (black bars) and flowers (white bars). Species names in bold indicate that flowers were collected and analysed for cardenolides. Sample sizes can be found in Table 1. Below the graphs is a molecular phylogeny of the 12 focal Asclepias species based on data from Agrawal & Fishbein 2008, where posterior probabilities can also be found.
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Table 1. The number of individual cardenolides found in nectar and leaf samples from the 12 focal Asclepias spp., along with number of compounds that are found in both plant parts
|Species||Nectar cardenolides (n)||Leaf cardenolides (n)||Shared cardenolides|
|A. angustifolia||0 (5)||0 (2)||N/A|
|A. barjoniifolia||4 (3)||5 (2)||3|
|A. boliviensis||5 (6)||7 (2)||3|
|A. candida||4 (3)||7 (2)||3|
|A. curassavica||0 (8)||11 (2)||N/A|
|A. fascicularis||0 (7)||0 (1)||N/A|
|A. incarnata ssp. pulchra||0 (2)||1 (1)||N/A|
|A. mexicana||1 (2)||1 (2)||0|
|A. nivea||11 (7)||13 (2)||9|
|A. perennis||9 (4)||16 (2)||8|
|A. pumila||7 (1)||6 (2)||4|
|A. texana||0 (4)||2 (3)||N/A|
Cardenolide sampling and quantification
We collected samples to analyse constitutive cardenolide concentrations on five occasions between July 2007 and August 2008. The frequency and timing of nectar collections depended on the flowering time of the different species. We collected nectar from all plants in flower, making collections one to three times per species over the course of the sampling period. We also collected whole mature leaves from all 12 species. Leaves were collected after taking nectar samples to avoid potential induction of cardenolides in nectar or leaves as a result of the leaf removal. After completing nectar sampling, we collected whole flowers in six of the 12 species. We were unable to collect flowers from all species because although nectar collection rarely resulted in damage that affected floral nectar quality, manipulation of flowers often led to petal damage that may have altered cardenolides in floral tissue.
We primarily collected nectar from flowers using 5-μL graduated microcapillary tubes, although several species (Asclepias angustifolia, A. curassavica and A. nivea) produced nectar so copiously that we used a 200-μL microcapillary tube for sample collection. We took every precaution to ensure that we caused no damage to the nectary, as this could induce cardenolide production; fortunately, damage can be visually detected in Asclepias because of the exudation of latex, which is rich in cardenolides (Zalucki, Brower & Alonso 2001). On the few occasions where damage did occur, samples were discarded. Because most plants produce very little nectar per flower, we pooled nectar; samples were pooled daily within individual plants, with each sample containing between 20 and 230 μL of nectar, representing tens to hundreds of individual flowers. Nectar was added to 500 μL of 70% ethanol (following Blüthgen, Gottsberger & Fiedler 2004) and stored at −80 °C prior to analysis. After nectar samples were pooled, we had between one and eight samples per species from 1 to 2 individual plants each. We then harvested leaves and flowers from a subset of plants also used for nectar sampling (for a total of 1 to 3 leaf and flower samples per species, collected as single leaves or umbels, respectively) and froze them immediately at −80 °C. Because our nectar samples typically lack independence (although temporally separated, they were often collected from the same plants), they cannot be considered true replicates. Therefore, although we analysed each sample separately, the data were combined to create composite profiles of cardenolide identities and concentrations for each of the 12 focal species.
We used high-performance liquid chromatography (HPLC) to quantify cardenolides in nectar, leaf and flower samples, adapting a protocol from Zehnder & Hunter (2007). We prepared nectar samples for extraction by drying down all water and ethanol from the stored samples using a rotary evaporator (Labconco, Kansas City, MO, USA). We extracted the residuum with 1 mL of 100% methanol and added 10 μL of a 0·2 g/L solution of the cardenolide digitoxin as an internal standard, which allowed for the direct comparison of unknown cardenolide peaks to a known cardenolide of predetermined quantity. Digitoxin is a common standard used when quantifying cardenolides using chromatography and is not known to be produced by Asclepias. We let the samples shake gently for 24 h and then centrifuged them for 30 min at 19722 rcf and 15 °C, removing the supernatant containing the dissolved cardenolides for further analysis. In some cases, preliminary analysis of nectar samples had very low cardenolide yield, so where necessary, we pooled material yet again to increase our power to detect cardenolides. For leaf and flower samples, we ground fresh tissue in liquid nitrogen and added 10 μL of 2 g/L digitoxin to each sample, at c. 100 mg of tissue (fresh mass). We extracted the samples with 1 mL of 100% methanol using FastPrep® homogenization (MP Biomedicals, Solon, OH, USA) to rapidly lyse cells set at 5·0 m/s for 30 s. Flower and leaf samples were spun down as with nectar samples, but we did not further concentrate the material.
Cardenolides were analysed on an Agilent 1100 series HPLC using a Gemini C18 reversed-phase column (3 mm, 150 × 4·6 mm; Phenomenex, Torrance, CA, USA). We injected 15 mL of extract, which was eluted at a constant flow rate of 0·7 mL/min with a solvent gradient of 0·25% phosphoric acid in water and acetonitrile as follows: 0–5 min with 20% acetonitrile, followed by a constant increase to 70% acetonitrile until 20 min; steady elution for 20–25 min with 70% acetonitrile; followed by a constant increase to 95% acetonitrile until 30 min; and hold at 95% acetonitrile until 35 min. UV absorption spectra were recorded from 200 to 400 nm and cardenolides were quantified by integrating the peak area at 218 nm. Each HPLC run included several blanks, containing only methanol and controls, comprising of methanol plus a known concentration of digitoxin. Cardenolides are identifiable from other compounds by a single symmetrical peak that absorbs between 217 and 222 nm (Zehnder & Hunter 2007; Rasmann, Johnson & Agrawal 2009), and we confirmed this by checking the shape and absorption of the peak in control samples containing only digitoxin. The amounts of cardenolides present were then calculated relative to the peak area of the digitoxin internal standard.
Because the concentrations of cardenolides in nectar are often very low, true peaks can be difficult to distinguish from noise. Therefore, we defined a detection threshold and considered all peaks with uncorrected peak areas of more than 15 absorbance units, representing >50 ng of cardenolides, as true peaks. Peaks that fell below this threshold were only considered true peaks if they met the following conditions: (i) the peak shape was extremely symmetrical, and (ii) a peak had been detected at that retention time in the nectar of at least two other species. Final cardenolide concentration estimates were calculated as nanograms of cardenolides per microlitre of nectar or per microgram of fresh tissue collected.
Large-bodied Lepidoptera and Hymenoptera, such as monarch butterflies, honeybees and bumblebees, are effective pollen vectors for most milkweed species (Woodson 1954; Wyatt & Broyles 1994). We used a commercially supplied Bombus impatiens (Biobest Canada, Leamington, ON, USA) colony for our behaviour experiments. We created an artificial nectar solution by mixing 30% w/w sucrose with the cardenolide digoxin (92% HPLC grade; Sigma, St. Louis, MO USA). Ideally, we would employ cardenolides from Asclepias for these experiments, but this was not feasible because of limited availability of plant tissue and the prohibitive costs associated with purifying cardenolides. We therefore used digoxin, an affordable, commercially produced cardenolide, as a proxy for Asclepias nectar cardenolides. Digoxin, found in Digitalis spp., causes 50% mortality in honeybees when ingested at concentrations of 0·5% or approximately 5 ng/μL (nectar consumed ad libitum for 48 h), but did not deter honeybees at concentrations of 1% or c. 10 ng/μL (Detzel & Wink 1993). Digoxin differs from digitoxin, the standard used in HPLC analyses, in that it is a more polar compound and is therefore more representative of the cardenolides found in Asclepias nectar.
Preliminary studies showed that bumblebees did not find digoxin deterrent at concentrations of 10 or 50 ng/μL (J. S. Manson, unpublished data). We therefore prepared three solutions for testing pollinator response to cardenolides in nectar: 100, 250 and 1000 ng/μL digoxin. We mixed nectar solutions every 2 days, refrigerating unused portions at 4 °C for no more than 24 h.
We examined pollinator behaviour using methods reported by Gegear, Manson & Thomson (2007). In short, marked worker bees were trained to associate artificial flower colour (either blue or yellow) with one of two nectar types, either artificial nectar composed of 30% sucrose or containing both sucrose and digoxin. Artificial flowers were constructed from microcentrifuge tubes with the caps removed and spray-painted polystyrene squares measuring c. 3 × 3 cm. Bees foraged freely as a group on alternating monotypic training arrays of each flower type, but experiments were conducted with single foragers. The association between flower colour and nectar condition was randomized among bees to control for any potential bias because of innate colour preferences. Immediately following training, individual bees foraged on a mixed array with 30 flowers of each type for at least 80 flower visits. We filled flowers with 2 μL of nectar and refilled each flower immediately after it was drained during the trial. After an individual worker had completed the minimum visit number, she was captured and terminated. We replaced flowers between individual bees to remove any scent marks left on artificial flowers after foraging, which might deter nectar collection. We evaluated a total of 24 individual foragers at three digoxin concentrations: 100 ng/μL (n = 10), 250 ng/μL (n = 8) and 1000 ng/μL (n = 6). All foraging bouts were videotaped and subsequently analysed using JWatcher Video Version 1.0 (Blumstein & Daniel 2007).
We assessed the effect of nectar cardenolides on two behaviour parameters, pollinator preference and flower-handling proficiency. We quantified preference by counting the number of visits to flowers with cardenolide-enriched and cardenolide-free nectar. A visit was defined as an event where a bee entered and imbibed nectar from an artificial flower. To evaluate the effect of nectar cardenolides on flower-handling proficiency, we identified bees with a significant preference for either nectar cardenolides or control nectar, as determined by G-tests and examined a series of sequential visits from these individual bees, calculating both the average visit duration and the rate of flower visitation.
We analysed quantitative differences in cardenolides between plant parts by summing the concentrations of all individual cardenolides within each sample, then averaging this summed total across all within-species samples for each plant part, creating an average gross cardenolide concentration (ng/μg for leaves and flowers or ng/μL for nectar). Sampling methods differed for each plant part. Leaf and flower samples were only collected from plants in flower and were taken either from two different plants on the same date or at two collection dates months apart. To minimize pooling of nectar, so as to have as many opportunities as possible to capture cardenolide diversity, nectar samples were often taken from different inflorescences on the same plant or from the same plant during different flowering periods. Despite this lack of independence, using an average over all samples provides the most representative quantitative data for total nectar cardenolides. Quantitative data are therefore reported and analysed as means per species, with variances calculated across all available samples from that species.
Different samples often varied in the identity and concentration of individual compounds. Cardenolide peaks were considered to represent the same compound when their retention times were within 0·1 min of each other. For our primary qualitative analysis, we created ‘cardenolide profiles’ for the nectar and leaves of each of the 12 species examined. Due to the limited number of species we were able to sample, we omitted flowers from these analyses. We created these profiles by pooling all within-species samples of nectar and leaves. We then summed the mass of each individual cardenolide across those samples and divide each sum by the total cardenolides within the samples, calculating the proportion that each individual cardenolide comprises for the total nectar or leaf cardenolides of each species (See Supporting information). The cardenolide profiles then locate each species' nectar and foliage in a multidimensional space, such that plant parts with similar profiles are close together.