Adaptation to changing conditions is one of the strategies plants use to survive climate change. Here, we ask whether plants’ leaf morphological and physiological traits/gas exchange variables have changed in response to recent, anthropogenic climate change. We grew seedlings from resurrected historic seeds from ex-situ seed banks and paired modern seeds in a common-garden experiment. Species pairs were collected from regions that had undergone differing levels of climate change using an emerging framework – Climate Contrast Resurrection Ecology, allowing us to hypothesise that regions with greater changes in climate (including temperature, precipitation, climate variability and climatic extremes) there would be greater trait responses in leaf morphology and physiology over time. Our found that in regions where there were greater changes in climate, there were greater changes in average leaf area, leaf margin complexity, leaf thickness and leaf intrinsic water use efficiency. Changes in leaf roundness, photosynthetic rate, stomatal density and the leaf economic strategy of our species were not correlated with changes in the climate. Our results show that leaves do have the ability to respond to changes in climate, however, there are greater inherited responses in morphological leaf traits than in physiological traits/variables, and greater responses to extreme measures of climate than gradual changes in climatic means. It is vital for accurate predictions of species’ responses to impending climate change to ensure that future climate change ecology studies utilise knowledge about the difference in both leaf trait and gas exchange responses, and the climate variables that they respond to.

Historic seeds were acquired for 32 species from stored collections in ex-situ seed banks at The Australian PlantBank and the Australian National Botanic Garden. This included four herbaceous species, ten shrubs, seven shrub-trees and eleven trees where all shrubs, shrub-trees and trees were evergreen species (See Everingham et al 2021, Ecology and Dryad dataset https://doi.org/10.5061/dryad.4f4qrfj83 for more information of seed collection). Matched modern seeds from the same species as the historic seeds were collected in the same location, at the same time of year as their historic counterparts. The amount of time between the historic and modern seed collections ranged from 29 years to 40 years.

Seeds were germinated on water agar (0.7% w.v.) in controlled incubators. Most species were germinated at 20°C with a 12-hour light, 12-hour dark cycle, but some species required specific germination treatments such as gibberellic acid (GA₃), smoke water (1%) or specific temperature and light treatments (see Everingham et al 2021, Ecology and Dryad dataset https://doi.org/10.5061/dryad.4f4qrfj83 for full germination treatment methods). Treatments were always kept constant for modern and historic seeds of each species. After germination, we transferred up to 50 germinated seeds to trays made up of 24-cells each measuring 4 cm (depth) by 2 cm² (square area) cells. The seedlings grew for two weeks in the trays to ensure early seedling survival before being transferred to individual 1.9 L pots. Potting soil comprised of 33% Australian Native Landscape supply of “Organic Garden Mix”, 33% washed river sand and 33% Cocopeat as well as a general slow-release fertiliser added at 200 mL per 75 L of soil. Plants were grown in a glasshouse at UNSW, Sydney for six months with an overhead irrigation system. Pots were randomised each month to reduce position effects.

After the six-month growing period, we measured a range of morphological leaf traits including leaf area, leaf roundness, leaf margin complexity and leaf thickness following standard protocols from Perez-Harguindeguy 2013, Australian Journal of Botany. 

To measure leaf shape, leaf area and leaf mass per unit area (LMA), we collected three fresh leaves (excluding the petiole) from each individual plant at the end of the six-month growing period. For two species (Acacia georgensis and Acacia concurrens), due to their seedling size, we were not able to measure area on three leaves and one to two leaves were sampled. Images of these fresh leaves were captured on a Flatbed Scanner and their area and shape metrics were calculated using values measured in image analysis software, ImageJ. Leaf surface area was calculated as the average of the three leaves’ total surface area.  ImageJ provided a measurement for each leaf of the maximum length (longest axis of the smallest possible rectangle drawn around the leaf) and width (longest axis perpendicular to the determined maximum length). From these measurements we calculated leaf roundness as the average ratio of width to length of the three leaves whereby the leaves with roundness measurements closer to zero would be longer, thinner leaves and the leaves closer or equal to 1 would be rounder leaves.  We calculated the margin complexity as the average of the ratio of perimeter length (cm) to surface area (cm2) from the perimeter of the leaf and the area analysed in ImageJ. To calculate leaf mass per unit area we used the leaf surface area calculations measured in ImageJ. The leaves were then dried to a constant temperature using a drying oven at 60° C for 72 hr. Oven dry mass (g) for the leaves was measured by weighing on a microbalance (Mettler Toledo© AG204 microbalance, 1 x 10-4 accuracy). LMA was calculated as oven-dry mass divided by fresh area. We measured leaf thickness by sampling one leaf from each individual modern and historic plant from all species (the third leaf from the growing tip, counted from the first fully developed/unfolded leaf). On these leaves we measured fresh leaf thickness (mm) at two points on adjacent sides of the mid-vein using a micrometer. An average for leaf thickness was taken from the two measurements for each individual plant. Finally, we calculated stomatal density using the clear nail polish peel method. Clear nail polish peels were performed on the first mature leaf closest to the growing apical tip from each plant. Clear nail polish was painted on the top and underside of the leaf on fresh tissue, away from the mid-vein or any prominent veins. We allowed the nail polish to dry for approximately 60 seconds before removing and mounting on a microscope slide with a coverslip. The peels were then imaged using a Leica© microscope. Stomata in each image were counted manually for the top of the leaf and the bottom of the leaf and the average stomatal density (stomata.cm-2) was calculated for each plant and use in further analysis.

We measured physiological variables including leaf photosynthetic rate, intrinsic water use efficiency (iWUE) and leaf nitrogen content. To obtain photosynthetic measurements, we used portable infrared gas analysers (LICOR 6400XT, Lincoln, Nebraska) on well-watered, non-root-bound, non-flowering individuals. We randomly selected a subset of ten historic plants and ten modern plants from each species. Some species had fewer than ten plants available, and some species were excluded from photosynthetic measurements because their leaves were not large enough to fit into the gas chamber without damage to the majority of the seedling. We took infrared gas measurements on the youngest fully expanded mature leaf following standard protocols [66] between the hours of 10:00 to 14:00 (Australian Eastern Standard Time) on days with no visible cloud cover. We ensured that for each species, infrared gas exchange measurements were taken on historic and modern plants at random within a 30-minute period to minimise changes in light or temperature. Our measurements were made under constant saturating light conditions (1800 μmol m^-2 s^-1) provided from a constant light source in the LICOR chamber. The chamber CO₂ concentration was set at 400ppm and the temperature set at 25° C. We took five consecutive measurements approximately two seconds apart and used the average of these five measurements. We recorded the light-saturated photosynthetic rate (A_sat; μmol CO2 m^-2 s^-1) and the stomatal conductance (g_s; mol H₂O m^-2 s^-1), and then calculated the intrinsic water use efficiency (iWUE) as the ratio between photosynthetic rate and stomatal conductance.

To quantify leaf nitrogen, we harvested leaves at six months, dried them for 72 hr at 60°C, pooled and homogenised each species’ individual modern leaves and individual historic leaves separately and then ground the dried leaf tissue. For each species we sent a pooled sample of historic ground leaf tissue and a pooled sample of modern ground leaf tissue to the Environmental Analysis Laboratory at Southern Cross University, Lismore, Australia for nitrogen analysis.

Climate change metrics were determined for each species’ historic and modern seed collection based geographically on modern seed collection site location data (which was collected typically at the same location as the historic data or within a 1 km radius) and were obtained from the Australian Gridded Climate Data at 5 km² resolution following methods from Everingham et al. 2021, Ecology. The processing code is freely available at https://github.com/SEveringham/ClimateData. The amount of change in all climate metrics was calculated across the five years before historic and modern seed collection to capture longer-term climate change responses of the species without extending to a period of climate that may become non-meaningful or overlap with modern climate data. The amount of change in precipitation metrics and heatwave duration were calculated using the log-transformed ratio of means. Change in all temperature metrics was calculated as the difference between the modern and historic climate metrics. We used different scaling methods because a difference of a few degrees Celsius of temperature has a much higher biological impact than a difference of a few millimetres of precipitation as precipitation has a much larger range of measurement than temperature. None of the climate change metrics were significantly correlated with one another (as all correlation coefficients were below 0.6) and therefore no climate metrics were excluded from our analyses.

The climate change metrics we used included the change between the modern and historic seed collections in mean monthly temperature (calculated as the daily median temperature in the month prior to the seed collection and averaged across the previous five years before the seed collection was made) and mean monthly precipitation (an average of precipitation from the month prior to seed collection and then averaged across the 5 years prior to collection). Both the change in the range of temperature and the range of precipitation were calculated as the change (between historic to modern collections) in the difference between the yearly maximum and minimum temperature or precipitation averaged across the five years prior to each seed collection. We also used metrics for change in temperature variability and change in precipitation variability, both of which were calculated as the coefficient of variation (standard deviation divided by the mean) of the temperature or precipitation of the month prior to seed collection averaged across the five years prior. The change in maximum and minimum precipitation of the season before collection were calculated to determine the effects of seasonal rainfall and these were an average across five prior years of collection of the maximum rainfall in the 4 months prior to seed collection (bound by wet season in the subtropics or autumn, winter, spring, summer seasons in the mid-latitudes). We used the change in vapour pressure deficit (VPD) as an indication of the change in atmospheric aridity between the historic and modern seed collections. Finally, metrics of change in extreme climate events included the calculation of maximum heatwave duration (the longest heatwave across all seasons in the 5 years prior to collection whereby heatwaves were defined based on exceptionally high air temperature following the relative extreme heat index metric) and maximum dry spell duration (following the same protocol as maximum heatwave duration but instead with dry spells as calculated from an “extreme dryness index” using VPD measurements).

All of the above raw data is available in the leaf measurement file and the climate variable file.

We performed all data transformation analysis in R, version 3.6.0 with code freely available at https://github.com/SEveringham/leaf-trait-responses-to-climate-change.

All transformed data is available in the full leaf analysis data file provided.

Change in traits or gas exchange variables was calculated for all morphological, photosynthetic and leaf economic traits or variables using the log-transformed ratio of means per species using the escalc function in the metafor package.

To determine if leaf economic spectra were related to changes in climate, we used a Principal Components Analysis (PCA) to obtain metrics that combined the change in inverse LMA, photosynthetic rate and nitrogen content. The inverse of LMA (specific leaf area [SLA]) was used as it is negatively related to leaf economy (i.e. leaves that have a larger surface area per unit mass will have a lower LMA and are typically on the ‘faster' end of the leaf economic spectrum). The PCA was achieved using the prcomp function in base R and used imputed data as not all species had measurements for all three variables (imputation was done using the imputePCA function in the missMDA package).

Data files can be opened in microsoft excel or any program that can read xlsx files