Explore the videos below to learn more about our research efforts, technologies and facilities.

Novel application of lasers in plant science research at Penn State University

Developed in the Roots Lab at Penn State University, laser ablation tomography is a novel method that allows for rapid, three-dimensional quantitative and qualitative analysis of plant anatomy.

Hi everyone. Welcome to the laser ablation tomography facility that we have housed here in the Department of Plant Science at Penn State University. Behind me you can see the latest iteration of a tool that we use for visualizing plant anatomy. It’s a really interesting technique that generates a lot of interest whenever we present data generated from this methodology at meetings or in academic journals. We get a lot of questions about what it is, how it functions, and the capacities that it can be used for. So to address all of these questions, I thought it would be easiest to summarize all of this in a video that shows you how the system works and how it can be used.

 Our group is most interested in visualizing both the root and shoot anatomy of major crops like bean, maize, and rice. We are looking at different anatomical features, specifically in the root system, that can help these major crops in both acquiring and more efficiently utilizing important resources like water, especially under drought scenarios, as well as important macronutrients like nitrogen and phosphorus, which can often be limiting in many agricultural soils. 

 Here we can see a general schematic of the laser ablation tomography system. Its base is a UV laser source which is projected at 355 nm, and it occurs at a pulsed repetition rate somewhere between 25 kHz and 40 kHz, which is appropriate for most of our samples. This laser is directed through some beam shaping optics, and then into a galvanometer which is used to oscillate the beam over a linear distance to create a cutting sheet. We then have a biological sample that is moved into the path of the beam by a motorized stage, and as the sample is ablated by the beam, a digital camera that is fitted with a macro lens is imaging each illuminated slice in real time.

 Here we can see the laser source, with the beam path being directed by a series of mirrors and shielded from the user by these opaque tubes. The metallic box is the galvanometer which rapidly oscillates the beam back and forth to create a cutting sheet that ablates the sample positioned below at the focal plane of the camera.

 Another feature of the laser ablation tomography system is its capacity for differentiation of spectra from different tissues under UV excitation. Red, green, and blue channels can be measured from tissues illuminated by the UV laser, and dominant emission wavelength can then be determined from these data to provide some information on the composition of the tissue. For example, here we can differentiate between the chitinous cell wall of arbuscular mycorrhizae colonizing this maize root sample, which is highlighted in red, and the auto-fluorescent spectra emitted from the lignin and cellulose based walls of the plant tissue. 

Laser ablation tomography also allows for three-dimensional visualization and quantification of features in samples. Image stacks of a sample can be compiled into 3D reconstructions, similar to Z stacks in confocal microscopy. While Z stacks generated through confocal techniques are ideal for cellular level visualization, the depth that’s able to be visualized using traditional microscopy is limited by the opacity of the tissue. Although laser ablation tomography does not have the same resolution of a microscope, it is ideal for tissue level visualization on the scale from 0.1 mm to about 1 cm, and three-dimensional reconstruction of large opaque tissue samples is possible.

Studying Drought Stress in the Field at Penn State University

Here we provide an overview of our research on drought at Penn State University's Russell E. Larson Agricultural Research Center, including a brief explanation of some of methods used in our work.

Hi, welcome to the Russell E. Larson Agricultural Research Center out here at Rock Springs, Pennsylvania. We’re just a couple miles from Penn State University’s main campus, and this is the site where we perform a lot of our field-based studies to investigate some of the physiological adaptations in root and shoot tissue that help major field crops like corn and bean perform well under drought scenarios, as well as soils that are limiting in both nitrogen and phosphorus. 

Behind me here you can see some of these specially managed fields we have to mimic a terminal or intermittent drought scenario. These are what we refer to as “rainout shelters”, and they function as large greenhouses that can move over our experimental plots along these rails every time a rain event occurs. Then as that rain event terminates, these shelters will move back off of the plots and allow normal climatic conditions to exist above these fields. So, we get normal evapotranspiration rates occurring when these structures are off the plots, but we can move them over to inhibit rainfall.

The movement of these rainout shelters is automated by rain sensors, programmed to move the shelter over the field when they detect rainfall and off the field when it dries. 

To help us monitor soil moisture throughout our fields as we impose drought and as drought progresses throughout our field trials, we utilize what are called time-domain reflectometry probes or TDR probes. These function by hooking them into a computer system with software that sends an electrical pulse down these cables and into probes we have buried at various depths throughout the field, and the resistance of that waveform that goes down this cable and into the soil tells us something about soil moisture. Additionally, we use other methods like gravimetric soil moisture that we collect using these types of cores, where we can collect physical samples and bring them back to the lab and get information on gravimetric water content by weighing the fresh sample, drying that soil sample out, and then weighing it again to see how much water was in the original sample when it was freshly collected.

Estimates of aboveground shoot biomass can be determined from stem diameter, taken at the base of the shoot just above the emerging brace roots. For this measurement, we can use a caliper.

Similarly, record of plant height also correlates strongly with aboveground biomass.

Time of flowering and the anthesis silking interval can have significant effects on yield under drought stress. Consequently, diligent record of days to flowering and the time between pollen shed and silk emergence for all varieties being evaluated, is essential

To help us measure parameters like carbon assimilation, transpiration, stomatal conductance, and leaf temperature, we can utilize this instrument here, known as the Licor-6800.

To help us better understand the mechanisms of drought adaptation that may exist among the different varieties we are evaluating in our drought trials, we can utilize metrics like leaf relative water content. For this metric, we would take leaf punches throughout the newest, fully expanded leaf in the canopy of a given variety. We would take several of these punches and store them in these airtight containers, then take these back to the lab to get the fresh weight of these discs. We would then suspend these discs in deionized water so they can absorb additional water they may have expelled through transpiration, and then we will dry and re-weigh these samples so we will get the fresh weight, the turgid weight and dry weight of these samples. 

How efficiently different varieties utilize the water they take up through their root system is largely dependent upon the anatomy of their leaves. We collect leaf anatomy samples for evaluation back at the lab to explore different components like stomatal density and stomatal size on the surface of the leaf, as well as a lot of the anatomical features that can be visualized within the leaf. Leaf samples collected for anatomical analysis are cut to include part of the midrib as well as the leaf lamina and are preserved in 75% ethanol.

To help us understand the degree of stress that plants are experiencing in our drought trials, we utilize this tool which is known as a pressure bomb or Scholander bomb. To understand how this device works, you need to know something about how water is transported through plants. So just a brief overview, plants are acquiring water from the soil through their roots, and they are transporting that water through a series of hollow cells, or tubes, known as xylem vessels. That water is transported in a continuous column of these xylem vessels from the roots, to the stems, and into the leaves where that water is then transpired out of the stomates during photosynthesis. As water is transpired out of these leaves during photosynthesis, water needs to be continuously pulled out of the soil, through these xylem vessels in the roots, the stem, and the leaves. This negative tension that exists throughout the entire plant can be measured using this device. Under drought stress where there is less available water in the soil, that negative tension throughout the entire plant where they are trying to pull up that water from the soil through the roots, stems, and leaves, is more severe than under more well-watered scenarios where there is more available water in the soil and less negative tension that exists throughout this column of water where plants can more easily transport that water from the soils, to the roots, to the shoots, to the leaves.

This device is able to measure that negative tension by taking a leaf sample and placing it in a pressurized chamber. What we are measuring is the amount of pressure that it takes for xylem sap to be extruded out of a cut surface of that leaf. The amount of pressure it takes for that sample to extrude sap out of the cut surface is equal and opposite to the amount of tension that exists throughout that plant. 

Under circumstances of drought stress, shallow soil horizons are the first to dry and often plant available water can still exist at depth even as drought progresses through the season. Therefore, crop varieties that have deeper distribution of their roots can access these water reserves later in the season and can often perform better in term of yield, compared to varieties that have a shallower distribution of their root length within the soil. To help us understand the genotypic differences that exist among different varieties in terms of their root distribution by depth, we utilize soil coring. Here we can place one of these soil coring tubes into the soil and extract a soil core that is contained in this metal tube. That soil core sample is then transferred to one of these sample holders and we will take this back to our lab at Penn State, divide this into increments by depth, and wash away the soil from the root length that has been captured within the soil core. This helps us to understand how different crop varieties have differences in root length distribution by depth, and then we can relate that to how well these different varieties perform under drought scenarios in the field. 

Since roots are directly responsible for soil resource capture, our group is particularly interest in genetic variation for root traits that may affect how efficiently different crop varieties acquire and utilize water under drought. To measure different components of root architecture and collect samples for anatomical analysis, roots must be excavated and washed in the field.

After soil is washed from the root crowns, shoots are separated from the roots and subsequently dried for measurement of aboveground biomass.

Here we can see some of the genotypic variation that exists for maize root system architecture. The root crown on the left has a higher degree of gravitropism in its nodal roots, which can provide a benefit under scenarios of drought stress by placing roots in deeper soil domains where water is more available. The root crown on the right has a lower degree of gravitropism and consequently will have a shallower distribution of roots and less access to water at depth.

Here we have a fresh maize root crown that was just excavated from the field, and we’ve taken it back to the lab to collect root samples. We will subsequently look at the anatomy of these root samples to identify genetic variation for anatomical traits that may affect how efficiently different varieties acquire and utilize water under drought. For example, genetic variation for anatomical traits like xylem vessels may affect how well different varieties transport water through the plant, and ultimately may impact yield under drought stress

Maize Root Anatomy Sampling

In this video we provide an overview of our methodology for collecting samples from the maize root crown for anatomical analysis.

Since roots are directly responsible for soil resource capture, our group is particularly interested in genetic variation for root anatomy that may affect how efficiently different crop varieties acquire and utilize resources like water, nitrogen, and phosphorus. To collect samples for anatomical analysis, roots must first be excavated and washed in the field.

Here we have a fresh maize root crown that was just excavated from the field, and we’ve taken it back to the lab to collect root samples for anatomical analysis. At first glance this maize root crown may look like one big tangle of roots, but upon closer examination we can see how the root system is organized into distinct root classes. 

Here we can see the mesocotyl, where the seed initially existed, and the primary root which emerged from that germinating seed. The rest of the subterranean roots you see in this crown are referred to as nodal roots and upon closer examination we can see they are organized into distinct rings or “whorls”. 

Here we see the first whorl of nodal roots. These are the oldest nodal roots in the crown, and are the first nodals to emerge after germination. As the plant goes through its vegetative stage of growth, new, subsequently thicker, nodal roots are emerging above the older roots in the crown. 

So here is the second whorl, third whorl, forth whorl, and then fifth whorl nodal roots. These pigmented roots, which were emerging above the soil surface, are called brace roots. 

Because these nodal roots are developing over time while stresses like drought may be becoming more severe through plant growth, our group is interested in collecting anatomy samples from across these different whorls to look at how the anatomy may change from whorl to whorl. 

A sharp set of pruners are an ideal tool for removing roots from the crown. It is important to be consistent in sampling location from crown to crown and we typically focus on the basal-most 2 cm of each nodal root, right where they emerge from the crown.

Anatomy samples collected will be stored in 2 ml tubes filled with 75% ethanol. This will preserve the root samples until we can image the anatomy of these roots using our laser ablation tomography system at a later date. Using our laser system, no additional prep work to fix or stain samples will be needed, and they can be sectioned directly from storage in the ethanol solution.

Genetic variation in the anatomy of these maize roots may affect the metabolic efficiency of the roots in foraging soil resources, the ability of roots to penetrate strong soils, or the transport of soil resources through the roots to the shoot. Anatomical features of interest from these samples include the size and abundance of cortical cells, xylem vessels, and air pockets in the cortex known as aerenchyma.