Team:Marburg/CFE Design

CFE Design

The goal of our project was to develop and optimise working cell free extracts of chloroplasts from different plant species. Cell-free systems serve as a prototyping platform for testing genetic parts, tracing metabolic pathways and screening potential antibiotics [1]. We have placed great emphasis on providing protocols for different end-users, so we selected the chassis based on agricultural impact and already existing scientific results.


For the decision on plants we decided on the model organisms Tobacco (Nicotiana tabacum), Spinach (Spinacia oleracea), Arabidopsis (Arabidopsis thaliana), as well as the single cell green alga Chlamydomonas reinhardtii. We also decided to focus on highly relevant crops, such as Maize (Zea mays B73), Wheat (Triticum aestivum) and Rice (Oryza sativa), as well as other plants that play a major role in world nutrition like Soybean (Glycine max), Canola (Brassica napus) and Tomato (Solanum lycopersicum).

For more Information on our choice of plants

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Chloroplast Cell-Free Systems

In the past, in vitro transcription or translation systems have been used in chloroplast basic research, even before chloroplast transformation was possible for any plant [2]. These systems allowed elucidating the foundational chloroplast biology, such as transcriptional [3] and translational regulation [4], light regulation of genes [5], their regulation via nuclear proteins [6] and their response to other environmental stimuli [7]. By utilizing high-throughput possibilities of cell-free technology and applying them to Plant Synthetic Biology via chloroplast cell-free systems, the time needed to test new genetic constructs can be hugely reduced. But at the beginning of our project only one protocol for the isolation of chloroplasts from Nicotiana tabacum was available. We wanted to expand the availability of protocols for the isolation of chloroplasts from other species.

Therefore we developed and optimized the protocols on growing and obtaining the used plant material, the isolation of intact chloroplast, chloroplast lysis, as well as the S30 extract preparation. In doing so, we oriented ourselves on a design-build-test-learn cycle for the extract preparation. Thus, we did not only base our optimization on our previous literature, but performed several rounds of experiment design.

More on the optimization of our protocols can be found

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Light and Temperature Conditions

Already when planting selected species, the growth conditions play a vehement role in the quality of the extracts. It is known that plants have different requirements regarding light and temperature conditions, for the stage of germination as well as for their growth in general.

At the beginning of our planting process in the greenhouse Tobacco, Wheat, Canola, as well as Arabidopsis thaliana were planted at the same time in the same conditions. This first round of planting served the purpose of investigating whether these plants, which normally follow a diurnal rhythm of 16 hours of light and 8 hours of darkness [8], can also grow in the lower light and temperature conditions of our greenhouse. Despite the fact that in optimal conditions tobacco should reach an adult stage with flowering after 4-6 weeks, the seeds germinated after 2 weeks and had grown only slightly after 4 weeks, while carrying yellowish leaves. This outcome was expected given that low temperatures can lead to differences in leaf color or even plant death [9].Similar observations were made on Arabidopsis and Canola.

With a device from biospherical instrument: QSL100, a spherical device to measure light intensities, we measured the light intensities of different locations inside the greenhouse and had to find out that the greenhouse only holds light intensities of about 55 µE. We consulted our PI for advice on the optimal light intensity for the plants. He recommended a minimum intensity of 150 µE for Solanaceae like Tobacco. Since it was certain that it would not be possible to constantly build up biomass with such a low growth rate, store-bought Spinach was used for isolation purposes throughout the winter. In order to generate biomass despite the poor conditions, we were given the opportunity to grow plants in phyto chambers at the University of Marburg. Thus, we were able to grow our first Tobacco plants in ideal light and temperature conditions with 120µE of light intensity and temperatures of 25°C by day and 18°C by night. In the beginning of spring we attempted our second try on planting a diverse set of plant species inside the greenhouse. This time our choices fell on Canola, another batch of Tobacco, Wheat as well as Rice, Arabidopsis and Cotton.


Since we decided to create cell-free systems of a variety of plants, their differences in germination conditions had to be evaluated. With the light conditions being around 120µE in the greenhouse as well, the Tobacco plants sprouted very fast, as well as Arabidopsis thaliana. However, we found that the biomass produced by Arabidopsis leaves is too little to generate the required 300g for chloroplast isolations in a steady repetitive manner, which led to our decision to not establish a cell-free system for this species.

During the germination time of the seeds we had to realize that Soybean, as well as Cotton and Rice did not germinate at all. To initiate germination, we decided to place the seeds in water in a germination jar and let them germinate in the phyto chamber at 100µE for 4 days, checking the germination status daily. After 4 days the sprouting of the Soybeans was clearly visible, whilst the Rice seeds did not show any changes. A slight swelling of the seeds of Cotton was also noticeable. The germinated seeds were transferred to pots filled with soil and placed in the greenhouse. Considering that the ideal temperatures for the growth of Oryza sativa are between 25°C and 30°C [10], we decided to adjust the water temperature for the germination of the seeds to that temperature and kept it at ~30°C. The water bath was put under direct sunlight since Rice is a light germ. After prolonged observation of 3 days, the swelling of the seeds was evident. The light germs were put onto heavily water drained soil and put into high light conditions, where germination proceeded further.

Despite the swelling of the seeds in Cotton, shoot formation did not occur. After further attempts with adjusted germination water temperatures of ~30°C, as well as the attempt by half-day spraying the covered seeds, placed 6cm into wet soil with distilled water for germination [11], we had to stop the generation of Cotton plants and decided to focus on the other plant species. For other light germinators such as Nicotiana tabacum and Arabidopsis thaliana, as well as Triticum aestivum, it was sufficient to achieve germination within a week by placing the seeds on the surface of moist soil. Canola being a light germ as well was placed 1 cm deep into wet soil with access to light. After 7 days, seedlings were present. No germination was necessary for Tomato, Spinach and oak since Tomato plantlets were bought, Spinach was bought freshly from the market and Oak leaves were harvested from the nearby forest.

Soil and Fertilization

In the greenhouse as well as in the phyto chambers, soil was used that had been mixed with clay. Before planting the phyto chambers, the soil was autoclaved. Initially, once a month a NPK 6-9-16 fertilizer high in potassium and phosphate was used for planting. After a few weeks, we noticed that the color intensity of the planted Canola and Tobacco plants had decreased. The leaves of Canola seemed to have turned slightly purplish and the Wheat was also visibly paler. It was decided to switch to a new fertilizer with a higher nitrogen content and to increase the frequency of fertilization to once a week. The newly selected NPK 16-16-12 fertilizer was chosen for its increased nitrogen content, since plant growth is dependent on an adequate supply of nitrogen in order to form the amino acids, proteins, nucleic acids, and other cellular constituents necessary for development [12].

After only two more weeks, a significant improvement in the coloration of the plants could be recognized, as well as faster growth, apart from the Canola plants. These showed only a slight improvement in condition. We hence decided to fertilize the Canola plants twice a week. Improvements in leaf condition were visible in the next generation of Canola plants. Since both Rice [13] and Tomato need potassium and phosphate for root and leaf growth, we fertilized them once a week with a NPK 20-20-20 fertilizer. Neither plant showed any deficit due to leaf coloration or leaf dieback.


With each planting, the seeds were placed in or on heavily moistened soil. Initially, all plants were watered daily or every two days. Daily watering was maintained for Tomato [14] and Rice [15], which need to be kept constantly moist. In the case of Rice, care was taken to keep the soil moist enough to form a small film of water on the surface of the soil. In addition, a constant humidity of 70% was maintained. Tomatoes were watered daily, but not as much as Rice. Care was taken to keep the soil moist, but the plants were not allowed to stand in water in the process.

Depending on weather conditions, Tobacco, Wheat, Soybean and Canola located in the greenhouse were watered at different intervals. In colder weather and low sunlight, rewatering was necessary only once a week. In high sunlight and temperatures up to 48°C in midsummer, daily watering became necessary. Thus, the rate of biomass production in the greenhouse was also strongly dependent on the seasons. By taking advantage of the constant environmental conditions within the phyto chambers, we were able to determine an optimum amount of water to add for the Tobacco plants growing there. With a diurnal rhythm of 16h light, 8h dark of 120 µE white light and temperatures of 25°C during the day and 18°C at night, 2.5 liters of water were added every three days to each tray containing 6 adult plants in pots. This value was determined by the average water consumption of the plants. For younger Tobacco plants, it was also important to keep the soil constantly moist. However, due to the fact that the younger plants consume less water, it was sufficient to add 1.5 liters of water to the trays every 3 days. Given that plants are expected to be in ideal conditions for chloroplast isolation from an age of 6 weeks [16], the rhythm of plant cultivation was optimized to meet this timeframe. Although we are certain that this optimal timeframe that is true for Tobacco plants can vary in others, this age at harvest seemed to work well for us with other plants too.

Chloroplast Isolation

The production of a cell-free system for chloroplasts includes 3 indispensable steps. Intact chloroplasts must first be isolated. These are then lysed and the lysate is further prepared. To isolate the chloroplasts, the specific plant mass must first be harvested, followed by homogenization of the leaf material, after which the chloroplasts must again be separated from the rest of the biomass, and thereafter only the intact chloroplasts are isolated.


For an optimal yield of intact chloroplasts it is inevitable to use actively growing, healthy plants. If previously harvested leaves are used, it is important to keep them in the cold and in the dark for no longer than one night before isolating the chloroplasts. After harvesting, the leaf material is washed and leaf veins are cut out. The leaf samples are then cut into small pieces to facilitate the subsequent homogenization step [17]. To reduce the enzyme activity of the leaf samples, the cut leaf pieces in an ice cold buffer are directly transferred to and kept on ice [18].

More Infos on dark incubation times here

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To homogenize the harvested leaf mass, we considered various methods. Blending the biomass with a homogenization buffer in a stainless steel blender is one option [19]. Here, the leaves are poured into a commercial blender with a leaf mass-buffer ratio of 1:3 and blended 3 times for 2 seconds at the highest level. The blended mass is filtered through 8 layers of muslin or fine nylon gauze to separate the remaining leaf material from the chloroplasts in the liquid. Instead of a series of muslin, Miracloth can also be used for filtering. Miracloth is an autoclavable filter material used for plastid isolation [20].

Another method of homogenizing leaf material from trees is the use of mortar and pestle. In this method, de-veined leaves are homogenized with pre-chilled mortar and pestle, in 2-4 ml pre-chilled isolation buffer by circular friction. This method is more precise and gentle, but effectively intended only for a small amount of leaf material [21]. Very fibrous leaves can also be ground with sand or metal beads and then blended or homogenized [22].

Similar to this method, and suitable for smaller amounts of leaf material, is the use of a filter cartridge. Here, 5-300mg of biomass is placed in the filter, a buffer is added and the filter contents are ground with a plastic rod with a twisting force. The substrate is then filtered by centrifugation [23]. Since we aim for a high yield of extract volume during production, but this method targets a small amount of biomass for isolation, this method is not suitable for our project.

Another alternative is the use of the enzymes cellulase and pectinase within leaf homogenization buffers. Cellulase catalyzes cellulose, which is an important component of the cell wall, to glucose, and. Pectinase degrades pectin middle lamellae. Both cellulose and pectin middle lamellae are important components of the plant cell wall. The use of these enzymes allows a gentle method of homogenization [24].

Finally, a homogenizer can also be used as an alternative. Using a homogenizer, the leaves are soaked in an isolation buffer and fractionated into beakers with another isolation buffer. The fractions are then homogenized with a homogenizer. [25] Similarly to the other methods, after using the homogenizer, a Miracloth is used for filtration. Another alternative for filtering is to use Cheesecloth, often used in combination with Miracloth.

Separation of Intact Chloroplasts

The next step during chloroplast isolation is the separation of the chloroplasts, intact as well as destroyed, from the rest of the leaf material. Low acceleration centrifugation makes it possible to perform this separation. Thereby, the centrifugation data in different papers on chloroplast isolation vary.
In a few isolations, no separation of chloroplasts have been performed, but after filtration the homogenate was put directly onto density gradients [26]. 4,000 rpm (2500 g) for 15 min are used for the chloroplast isolation of Marchantia polymorpha.
Meanwhile for the isolation of conifers the homogenate was centrifuged at 200 g for 15 min at 4°C. Since both species are less comparable to the species chosen for our project, we have not oriented ourselves according to these protocols.
The Chloroplast Isolation Kit from Sigma Aldrich has been tested for use with Spinach, Pea, Lettuce, Cabbage, Mangold, and Tobacco suggests the centrifugation for 7 minutes at 1,000 g to sediment the chloroplasts as a green pellet [27].
This separation is followed by another sequence of separation of intact chloroplasts from the destroyed plastids. The gradient centrifugation method is used to separate intact chloroplasts from the destroyed ones. Due to the size and density of chloroplasts, which vary in different species, diverse gradients are also prepared for different species. A continuous Percoll gradient is used for the separation of intact chloroplasts from Arabidopsis thaliana. [28] Also for Millet, a continuous percoll gradient of 10%-50% is normally used to purify the intact chloroplasts. [29]

Percoll is a classic medium for density gradient centrifugation of cells and subcellular particles. Centrifugation protocols using Percoll must be carried out in a swinging-bucket rotor. It is not recommended to use a fixed-angle rotor due to the intact chloroplasts smearing along the side of the tube. For Nicotiana tabacum, Percoll gradients of 20%, 50% and 80% are used [16]. For isolation of Tomato plant chloroplasts, Percoll is spiked with PEG-8000 and Ficoll-400 [30]. Since Percoll also serves as a good medium for bacterial and fungal growth, PEG8000 is added to inhibit microbial growth. Ficoll400, like Percoll, is a nonionic polymer for density gradient centrifugation [30]. As an alternative to Percoll, sucrose gradients with concentrations of 30% 40% and 60% are also used for the isolation of intact Spinach chloroplasts. Using sucrose instead of Percoll can drastically reduce the cost of the chloroplast isolation and make it more accessible for future iGEM teams.
Since our project focuses on a wide variety of plants, we used diverse concentrations and media for gradient centrifugation. After centrifugation, which takes place at various accelerations, usually between 10000g and 35000g, the intact chloroplasts form a band that is carefully removed and collected with a pasteur pipette.

More Infos about our low resource alternative

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Thereupon, the intact chloroplasts must be purified from the remnants of the substance used for gradient centrifugation to be prepared for lysis. Several rounds of centrifugation with the intact chloroplasts being mixed with a washing buffer at low accelerations are performed. Many washing buffers contain the chemical beta-mercaptoethanol, which destroys disulfide bridges and thus irreversibly inhibits RNases, thus preventing the degradation of RNA [32]. In most cases, the washing steps are repeated twice at a speed of about 2000g. The running time is usually 3-6 minutes [22] ,16].
Once the intact chloroplasts are freed from the gradient centrifugation media by centrifugation, they are resuspended in lysis buffer to prepare them for upcoming lysis. Most lysis buffers contain various salts, including magnesium and potassium acetates to maintain osmolarity and the ideal pH. These salts are added to the buffer with molarities between 50 and 150 mM [32].


To develop our cell-free systems from intact chloroplasts, the contents of the chloroplasts are freed from the organelle by disrupting its membranes. This process is called lysis. One method for the lysis of plastids is the use of a french press. Inside the french press, cells/organelles are passed through a narrow valve, subjecting the liquid to shear stress and decompression, resulting in cell rupture [33]. However, since lysis of chloroplasts with a french press is not yet an established method, this method was not further explored. The concept of needle lysis is similar to that of the french press. Sheer stress enacted when passing the chloroplast suspension through the needle causes rupture of the membranes. The decisive factor is not the diameter of the needle opening, but the number of times the extract is passed through the respective needle. For chloroplasts of Nicotiana tabacum, 25G needles are used, through which the intact chloroplasts mixed with lysis buffer are pressed through between 12 and 42 times [16].
This lysis method is time-consuming and requires uniform execution, but it is very cost-saving and easily available. In sonication lysis, the sonication probe generates ultrasound at a frequency greater than 20 kHz, which disrupts the cell and organelle walls. The chloroplasts are resuspended in a buffer and kept on ice to avoid overheating and denaturation. In plastids, sonication is usually performed with energies between 100J and 500J. Due to the limited availability of a sonicator at the University of Marburg, it was temporarily possible to attempt this variant of lysis.
In another lysis method, chloroplasts are ruptured by freezing and thawing. The osmotic shock induces rupture of the membranes [34]. For Pea, the chloroplast membranes are ruptured by a freeze-thaw cycle in which the suspension was placed in a -20°C freezer for 1 hr and then at room temperature until thawn. Although the method of shock freezing and subsequent thawing is easy to perform, it does not guarantee the total lysis of all chloroplast membranes. However, it is well applicable as an additional method to other lysis options.

For single cell green algae, lysis is chemically induced by the substances DEAE dextran and poly-D,L-lysines, which are polycationic macromolecules [35]. However, since we are working with vascular plants, we have refrained from this method. After performing the respective lysis methods, many protocols add an amino acid solution, as well as GTP, to maintain the activity of ribosomes of the lysed chloroplasts. After lysis, the interior of the chloroplasts is separated from the remaining destroyed membranes. High-speed centrifugation is used to settle membranes as sediment. Optionally, a dialysis can be performed after the lysis. The main goal of the dialysis of extracts is the removal of contaminants, but the process also causes changes in magnesium and potassium acetate concentrations. The supernatant produced after the preliminary centrifugation step is dialyzed against an S30 buffer for 3 hours with a Slide-A-Lyzer cassette [36].

Our Final Protocol

Despite minor variations along the way, we followed a main protocol for isolating chloroplasts.

The selected plants were incubated in darkness for 24 hours (Canola, Soybean), 48 hours (Tobacco, Wheat, Maize, Rice, Tomato) or 7 days (oak) prior to isolation, depending on the species. All the procedures of the actual chloroplast isolation and subsequent preparation of the extracts was carried out steadily in a cold room at 4°C.

Harvesting of the plant leaf material took place on the day of isolation. Leaf veins were excised from the selected leaves and 300g of leaf mass was used per isolation. The leaf material was washed extensively before further preparation. If leaf material was used from outside the greenhouse or phyto chamber, as in the case of Oak, it was cleaned with a bleach solution to remove possible contaminants and pathogens before further processing. The harvested leaf material was cut into small pieces and stored on ice.

For homogenization, we used a 1 liter commercial stainless steel Waring blender that was rinsed with desalinated water before each isolation. Again, depending on the species, certain leaf material homogenization ratios were used. 1 part leaf mass to 3 parts buffer was used for Tobacco, Canola, Soy and Spinach, while a ratio of 1: 4 was used for Wheat, Rice, Oak, Tomato and Maize. The mixture was blended 2 times for 5 seconds at the highest level and then again for 2 seconds. Should it become apparent that not all of the material has been blended properly, additional homogenization buffer can optionally be added in 100 ml steps. The following blending steps should be only 2 seconds each in order not to damage the chloroplasts excessively.

Prior to isolation, the filtration material, which is two layers of Miracloth on two layers Cheesecloth, was autoclaved. This is used to filter the blended leaf homogenate and collect it afterwards in a cold beaker. The filtrate is divided into 0.25 l polycarbonate bottles and directly stored on ice.

To collect the chloroplasts at the bottom of the bottles, centrifuge the filtrates at 1000g for 8 min at 1°C-4°C. Discard the supernatant and add 2 ml of the homogenization buffer to the sediment. Resuspension of the chloroplast pellet is done by carefully swinging the centrifugation bottles in a circular motion.

We used both Percoll and sucrose to form the density gradients in our project. Percoll does work better, but is also far more expensive than sucrose. For the separation of intact chloroplasts, Percoll gradients of 80%, 50% and 20% were prepared in 50 ml falcon tubes for all plant species except Spinach. For Spinach, sucrose gradients of 60%, 40% and 30% were used instead. The gradients were stored steadily on ice. Onto each prepared gradient tube, 2.5ml of chloroplast solution was carefully added.

Gradients were centrifuged at 10000g for 10 min at an acceleration and deceleration of 1 each in a fixed angle rotor. It is crucial to keep the acceleration and deceleration speed low in order to not disturb the gradients. Even though a swinging bucket rotor would be optimal, only a fixed angle rotor was available.

After centrifugation, two bands form in the gradient. The second band contains the intact chloroplasts. This band is carefully removed with a 25 ml glass pipette and transferred to new tubes, while not exceeding 12 ml of chloroplasts per tube. The falcon tubes are then filled up to 50 ml with a chloroplast washing buffer. These too remain steadily stored on ice.

Wash centrifugation was performed at 1000g for 4 minutes, whereupon the supernatant is discarded. The sediment is mixed with 3-5 ml of washing buffer per tube and resuspended. The solutions are collected and centrifuged again at 1000g for 4 min. Another time the supernatant is discarded. The sediment is resuspended in a total of 3 ml wash buffer and aliquoted into pre-weighed eppis standing on ice. These are centrifuged at 5000 rpm for 2 minutes in the final washing step. The supernatant is discarded carefully. Make sure to remove as much as possible!

Reweigh the filled eppis to determine the amount of chloroplast extract. Lysis buffer is added to the extract in equal amounts to the extract and is mixed by resuspension. After shock freezing in liquid nitrogen, the isolated chloroplasts can be stored at -80°C.

Frozen chloroplasts are on ice for 20 minutes prior to lysis. For lysis, chloroplasts are passed through 25G needles 24 times. Care must be taken to ensure that they are passed through steady speed! 1 μl of 0.1 mM GTP and 6 μl of 0.04mM amino acid solution per ml of extract are added and the suspension is mixed carefully.

Lysed chloroplasts are centrifuged twice for 30 minutes each at 30000g in an ultracentrifuge using tubes rated to withstand such forces. After the first round of centrifugation, the supernatant is carefully removed and transferred into a fresh eppi, which is then used for further centrifugation.

When aiming for expression with endogenous transcription, it is useful to perform dialysis after this centrifugation. Dialysis is performed by transferring the extracts into dialysis cassettes and running the dialysis for 4 hours in dialysis buffer. After 2 hours, the dialysis buffer should be renewed. This is followed by another centrifugation at 30000g for 20 minutes.

The resulting supernatant is flash frozen with liquid nitrogen and stored at -80°C. The finished cell-free system can be further used for measurements by thawing on ice for 20 minutes.

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