We thank Prof. Mark Abbe (UGA) for providing the NIR imaging equipment and Dr. Rasik Raythatha (Solvay Performance Chemicals) for the barium carbonate used in this work. We acknowledge the efforts of Isaiah Norris (UGA undergraduate) and Terra Blevins (North Oconee High School), who helped test the synthetic methods.

1. Preparation of CaCuSi4O10
<ol>
	<li>Melt Flux Synthesis of CaCuSi4O10
	<ol>
		<li>Weigh out CaCO3, SiO2, and Cu2CO3(OH)2 in a 2:8:1 molar ratio: 0.1331 g (1.330 mmol) of CaCO3, 0.3196 g (5.319 mmol) of SiO2, 0.1470 g (0.6648 mmol) of Cu2CO3(OH)2. In addition, weigh out the flux components (12.5% by weight): 0.0375 g of Na2CO3, 0.0125 g of NaCl, and 0.0250 g of Na2B4O7.10H2O. Add these materials to a clean agate mortar.</li>
		<li>Hand grind for ~5 min with an agate pestle until the mixture becomes a homogeneous light green powder (Figures 1a and 2a). Transfer this mixture to a clean, dry platinum crucible.</li>
		<li>Heat the crucible in a furnace to 875 &#176;C (ramp rate of 2 &#176;C/min), hold at 875 &#176;C for 16 hr, and then cool down to room temperature (rate of 0.8 &#176;C/min).</li>
		<li>Remove the crystals from the crucible and gently crush them using a pestle.</li>
		<li>Allow the crystals to soak in 50 ml of 1 M aqueous HCl overnight to remove the melt flux.</li>
		<li>Filter the crystals and wash with deionized water to fully remove any remaining melt flux. 
		Note: This material should be ground into a finer powder for powder X-ray diffraction (PXRD) analysis (Figure 5). It also can be characterized by optical microscopy (Figure 3), scanning electron microscopy (SEM) (Figure 4), and NIR photography (Figure 8).</li>
	</ol>
	</li>
	<li>Solid State Synthesis of CaCuSi4O10
	<ol>
		<li>Weigh out CaCO3, SiO2, and CuO in a 1:4:1 molar ratio: 0.1331 g (1.330 mmol) of CaCO3, 0.3196 g (5.319 mmol) of SiO2, and 0.1058 g CuO (1.330 mmol) and add to a clean agate mortar.</li>
		<li>Dampen the powder mixture with 1-2 ml acetone and hand grind with an agate pestle for ~5 min. Transfer the resulting light gray powder (Figures 1b and 2b) into a platinum crucible.</li>
		<li>Heat the crucible in a box furnace to 1,020 &#176;C at a ramp rate of 5 &#176;C/min, hold for 16 hr, and then cool down to room temperature</li>
		<li>Scrape out the loose, light blue-gray powder using a polytetrafluoroethylene (PTFE) spatula. 
		Note: The product can be characterized by optical microscopy (Figure 3), SEM (Figure 4), PXRD (Figure 5), and NIR photography (Figure 8).</li>
	</ol>
	</li>
</ol>
2. Synthesis of BaCuSi4O10
<ol>
	<li>Melt Flux Synthesis of BaCuSi4O10 
	<ol>
		<li>Weigh out BaCO3, SiO2, and CuO in a 1:4:1 molar ratio: 0.2085 g BaCO3 (1.057 mmol), 0.2539 g SiO2 (4.226 mmol), and 0.0840 g CuO (1.056 mmol). In addition, weigh out the flux component (12.5% by weight): 0.0765 g of PbO. Add these materials to a clean agate mortar.</li>
		<li>Hand grind for ~5 min with an agate pestle until the mixture becomes a homogeneous light gray powder (Figures 1c and 2c). Transfer this mixture to a clean, dry platinum crucible.</li>
		<li>Heat the crucible in a furnace to 950 &#176;C (ramp rate of 2 &#176;C/min), hold at 950 &#176;C for 24 hr, then slowly cool down to 700 &#176;C (rate of 0.1 &#176;C/min), and finally cool to room temperature.</li>
		<li>Remove the crystals from the crucible and gently crush them using a pestle.</li>
		<li>Allow the crystals to soak in 50 ml of 1 M aqueous HNO3 overnight to remove the melt flux.</li>
		<li>Filter the crystals and wash with deionized water to fully remove the remainder of the melt flux. Note: This material should be ground into a finer powder for PXRD analysis (Figure 6). It also can be characterized by optical microscopy (Figure 3) and NIR photography (Figure 8).</li>
	</ol>
	</li>
	<li>Solid State Synthesis of BaCuSi4O10
	<ol>
		<li>Weigh out BaCO3, SiO2, and CuO in a 1:4:1 molar ratio: 0.2085 g BaCO3 (1.057 mmol), 0.2539 g SiO2 (4.226 mmol), and 0.0840 g CuO (1.056 mmol) and add to a clean agate mortar.</li>
		<li>Dampen the powder mixture with 1-2 ml acetone and hand grind with an agate pestle for ~5 min. Transfer the resulting light gray powder (Figures 1d and 2d) into a platinum crucible.</li>
		<li>Heat the crucible in a box furnace to 960 &#176;C at a ramp rate of 5 &#176;C/min and hold for 16 hr, then cool to room temperature.</li>
		<li>Scrape out the loose blue powder using a polytetrafluoroethylene (PTFE) spatula. Note: The product can be characterized by optical microscopy (Figure 3), PXRD (Figure 6), and NIR photography (Figure 8).</li>
	</ol>
	</li>
</ol>
3. Exfoliation of CaCuSi4O10
<ol>
	<li>Charge a 50 ml round bottom flask with 0.50 g of CaCuSi4O10, 40 ml of deionized water, and a glass-coated magnetic stir bar.</li>
	<li>Attach a water-cooled condenser to the flask. Heat the reaction to 85 &#176;C with magnetic stirring at 400 rpm for two weeks.</li>
	<li>Remove from the heat source, allow the solution to settle undisturbed overnight, and then filter the supernatant through a 0.4 &#181;m membrane filter. Vacuum dry the solids. Note: The product is a light blue powder that can be characterized by optical microscopy (Figure 3), PXRD (Figure 5), transmission electron microscopy (TEM) (Figure 7), and NIR photography (Figure 8).</li>
</ol>
4. Exfoliation of BaCuSi4O10
<ol>
	<li>Charge a 50 ml plastic centrifuge tube with 0.14 g of BaCuSi4O10 and 20 ml of N-vinyl pyrrolidone.</li>
	<li>With the centrifuge tube immersed in an ice/water bath, sonicate with a probe ultrasonicator at 40% amplitude (17 W) for 1 hr.</li>
	<li>Let the dispersion settle undisturbed overnight, and then decant the supernatant into a new centrifuge tube.</li>
	<li>Spin down at 10,286 x g using a centrifuge. Decant the supernatant, leaving the nanosheets at the bottom of the centrifuge tube.</li>
	<li>Resuspend this material in 20 ml of water with a few minutes of bath sonication. To isolate a powder, filter through a 0.4 &#181;m membrane filter and vacuum dry the solids. Note: The product is a light blue powder that can be characterized by optical microscopy (Figure 3), PXRD (Figure 6), TEM (Figure 7), and NIR photography (Figure 8).</li>
</ol>
5. Ink Preparation
<ol>
	<li>Disperse ~0.10 g of CaCuSi4O10 nanosheets in 5 ml&#160;of deionized water using bath sonication for ~10 min. Note: This ink (Figure 9) can be used for painting, printing, etc. See Figure 10 for a representative example where the ink was applied to paper with a brush.</li>
</ol>
6. Near Infrared Photographic Imaging
<ol>
	<li>Irradiate the samples using red light (e.g.&#160;with a red light-emitting diode array), taking care to eliminate any other sources of light.</li>
	<li>Photograph using a camera modified to image in the near infrared region. Use f stop setting f/22 and an exposure time of 0.5 sec.</li>
</ol>

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The authors have no competing financial interests.

The preparation of Egyptian blue pigment, a mixture of mostly CaCuSi4O10 and SiO2, is a well-studied process4,13-21. The numerous reported procedures may be categorized as either melt flux or solid-state reactions. Two major advantages of the melt flux approach are that it permits lower reaction temperatures (&#60;900 &#176;C) and allows CaCuSi4O10 crystals to nucleate and grow from a molten glass phase20. The flux component is typically an alkali salt (e.g.&#160;Na2CO3) or borate compound (e.g.&#160;borax). In comparison, the solid-state syntheses omit the flux but require higher temperatures (~1,000 &#176;C) for the reaction between Ca, CuO, and SiO2 sources to reach completion.
Although the synthesis of Han blue pigment is not as well studied as that of Egyptian blue4,22-25, the preparation of BaCuSi4O10 follows similar melt flux and solid-state routes with two differences: (1) a PbO flux should be used, and (2) the reaction temperatures must be more closely controlled because of alternative Ba-Cu-Si-O phases that can form (e.g.&#160;BaCuSi2O6).
These points are illustrated by the detailed procedures and results described in this paper. First, for all methods, the starting materials should be ground to a smooth powder (Figures 1a-d) consisting of 5-20 &#181;m particles (characterized by SEM; Figures 2a-d). Next, the use of a significant amount of flux (12.5% by weight) in the preparation of CaCuSi4O10 and BaCuSi4O10 leads to highly crystalline products, which are characterized by intense blue coloration (Figures 3a and&#160;3c), relatively large particle sizes (Figure 4a), and strong PXRD patterns (Figures 5a and 6a). The diminished isolated yields (~70%) from these preparations are caused by adhesion of the melted reaction mixtures to the crucible. In comparison, CaCuSi4O10 and BaCuSi4O10 prepared by the solid-state route exhibit less intense coloration (Figures 3b and 3d) and smaller particle sizes (Figure 4b). As synthesized, these products are powders that can be isolated in near-quantitative yields. Thus, for both CaCuSi4O10 and BaCuSi4O10, the advantages of flux and the importance of reaction temperature cannot be overstated.
Remarkably, the exfoliation of CaCuSi4O10 and BaCuSi4O10 occurs under simple aqueous conditions. In the case of CaCuSi4O10, this reaction is quite slow at room temperature (&#8805;6 weeks to see any appreciable exfoliation), but it becomes synthetically useful at 80 &#176;C (substantial exfoliation after 2 weeks). In comparison, the exfoliation of BaCuSi4O10 is sluggish even at 80 &#176;C, and so we apply an even greater energy input in the form of ultrasonication. These reactions are highly reliable with two caveats. For CaCuSi4O10, it is important to use a glass-coated stir bar; if a standard PTFE-coated stir bar is used, we find that PTFE byproducts contaminate the CaCuSi4O10 nanosheet product. For BaCuSi4O10, it is important to control the ultrasonication power and time so that the reaction is stopped before the nanosheets become degraded.
Transmission electron microscopy (TEM) of the nanosheet products shows that these very thin materials have lateral dimensions ranging from hundreds of nanometers to several microns. In general these lateral dimensions correlate with the crystallite size of the three-dimensional starting material. In prior work, atomic force microscopy provided topographic mapping that demonstrated the single-layer thicknesses (~1.2 nm) of these nanosheets12. Photographs of powder CaCuSi4O10 and BaCuSi4O10 nanosheet samples (Figures 3e-h) show that their color is less intense than that of the starting materials, a direct result of nanostructuring.
Additional information is provided by PXRD (Figures 5 and 6), which reveals basal cleavage along the (001) plane and preferred orientation along the {00l} series for all nanosheet samples. These features reflect the stacked alignment of these highly anisotropic nanomaterials when drop-cast onto a substrate. Furthermore, the characteristic NIR emission of CaCuSi4O10 at ~910 nm and BaCuSi4O10 at ~950 nm is illustrated in a NIR photograph of all eight samples (Figure 8).
The solution processing of CaCuSi4O10 can be accomplished by simply preparing a colloidal dispersion of CaCuSi4O10 nanosheets (Figure 9) to use as an ink. This ink then can be applied to a substrate via spin coating, spray coating, ink jet printing12, or simply brushing (Figure 10). Importantly, the NIR emission properties of CaCuSi4O10 are retained at all stages of this process. These new possibilities highlight the contrast between CaCuSi4O10 nanosheets and the traditional use of Egyptian blue pigment, a highly granular material that is challenging to incorporate into a smooth paint.

Vibrant colors were prized throughout the ancient world. Even today, we can still see the remains of pigments and dyes created by every major culture. Remarkably, two of the most famous synthetic blue pigments share a similar chemical composition and structure, despite having been developed at widely different times and places. The colored components of both Egyptian blue, CaCuSi4O10, and Han blue, BaCuSi4O10, belong to the alkali earth copper tetrasilicate series, ACuSi4O10 (A = Ca, Sr, Ba)1, as well as the larger gillespite group, ABSi4O10 (B = Fe, Cu, Cr)2,3.
Beyond traditional pigment applications, current scientific interest in these materials centers on their strong near infrared (NIR) emission properties. This emission originates from the Cu2+ in square planar coordination; these ions are linked by tetrahedral silicate moieties within the three-dimensional crystal structure, and the resulting layers alternate with alkali earth ions4-6. Recent technical highlights include NIR imaging to identify Egyptian and Han blue pigments on cultural heritage artifacts7,8, lanthanide doping of ACuSi4O10 to enhance NIR reflectance properties and open new energy transfer pathways9,10, the use of ACuSi4O10 as the active material for optical sensors11, and the exfoliation of CaCuSi4O10 into monolayer nanosheets12.
In particular, this last example provides a way to nanostructure CaCuSi4O10 so that it can be handled as a colloidal dispersion rather than as a particulate solid12. Because colloidal dispersions are compatible with solution-processing techniques (e.g.&#160;spin coating, ink jet printing, layer-by-layer deposition), this advance opens new application areas that range from security inks to biomedical imaging. The experimental protocols illustrated in this contribution will enable researchers from diverse backgrounds to prepare, characterize, and use CaCuSi4O10 and BaCuSi4O10 nanosheets in their work.

<table><tbody><tr><td>Sodium carbonate (Na&lt;sub&gt;2&lt;/sub&gt;CO&lt;sub&gt;3&lt;/sub&gt;)</td><td>Sigma Aldrich</td><td>S7795</td><td>bioXtra, &amp;ge;99.0%</td></tr><tr><td>Calcium carbonate (CaCO&lt;sub&gt;3&lt;/sub&gt;)</td><td>Sigma Aldrich</td><td>C4830</td><td>bioXtra, &amp;ge;99.0%</td></tr><tr><td>Barium carbonate (BaCO&lt;sub&gt;3&lt;/sub&gt;)</td><td>Solvay Performance Chemicals</td><td></td><td>Research sample: Electronic-grade purity, nanocrystalline</td></tr><tr><td>Copper (II) carbonate basic [Cu&lt;sub&gt;2&lt;/sub&gt;CO&lt;sub&gt;3&lt;/sub&gt;(OH)&lt;sub&gt;2&lt;/sub&gt;]</td><td>Sigma Aldrich</td><td>207896</td><td>Reagent grade</td></tr><tr><td>Copper(II) oxide (CuO)&amp;nbsp;</td><td>Sigma Aldrich</td><td>450812</td><td>99.99% trace metals base</td></tr><tr><td>Silicon dioxide (SiO&lt;sub&gt;2&lt;/sub&gt;)</td><td>Sigma Aldrich</td><td>S5631</td><td>~99%, particle size 0.5-10 &amp;mu;m (approximately 80% between 1-5 &amp;mu;m)</td></tr><tr><td>Sodium tetraborate decahydrate (Na&lt;sub&gt;4&lt;/sub&gt;B&lt;sub&gt;4&lt;/sub&gt;O&lt;sub&gt;7&lt;/sub&gt;&lt;sup&gt;.&lt;/sup&gt;10H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Sigma Aldrich</td><td>S9640</td><td>ACS &amp;ge; 99.5%</td></tr><tr><td>Sodium chloride (NaCl)&amp;nbsp;</td><td>Sigma Aldrich</td><td>S9888</td><td>ACS &amp;ge; 99.0%</td></tr><tr><td>Lead(II) oxide (PbO)&amp;nbsp;</td><td>Sigma Aldrich</td><td>402982</td><td>ACS &amp;ge; 99.0%</td></tr><tr><td>N-Vinylpyrrolidinone (C&lt;sub&gt;6&lt;/sub&gt;H&lt;sub&gt;9&lt;/sub&gt;NO)</td><td>Sigma Aldrich</td><td>V3409</td><td>contains sodium hydroxide as inhibitor, ACS &amp;ge; 99.0%</td></tr><tr><td>Box Furnace</td><td>Thermo Scientific</td><td></td><td>Lindberg Blue M</td></tr><tr><td>Box Furnace</td><td>Carbolite</td><td>CWF 12/5 1200C</td><td></td></tr><tr><td>Bath Sonicator</td><td>Branson</td><td></td><td></td></tr><tr><td>Ultrasonicator</td><td>Qsonica</td><td></td><td>Misonix S-4000</td></tr><tr><td>Camera</td><td>custom modification of Nikon D3000 camera by LDP LLC MaxMax.com</td><td>n/a</td><td>Xnite Nikon D3000 camera with a Nikkor 18-200 mm lens and a Xnite 830 filter&amp;nbsp;</td></tr><tr><td>Light Source</td><td>Excled Ltd.</td><td>PAR64</td><td>LED Colour Beamer</td></tr><tr><td>Light Microscope</td><td>Leica</td><td></td><td>mz6 Stereomicroscope with Spot Idea camera and Software</td></tr><tr><td>Powder X-Ray Diffractometer</td><td>Bruker</td><td></td><td>D8-Advance diffractometer (Co-K&amp;alpha; radiation source)</td></tr><tr><td>Transmission Electron Microscope</td><td>FEI</td><td></td><td>Technai 20</td></tr><tr><td>Scanning Electron Microscope</td><td>FEI</td><td></td><td>Inspect F</td></tr><tr><td>Membrane filters</td><td>Millipore</td><td>HTTP04700</td><td>Isopore Membrane filter with 0.4 &amp;micro;m pore size</td></tr></tbody></table>

exfoliation of egyptian blue and han blue, two alkali earth copper silicate-based pigments

In a visualized example of the ancient past connecting with modern times, we describe the preparation and exfoliation of CaCuSi4O10 and BaCuSi4O10, the colored components of the historic Egyptian blue and Han blue pigments. The bulk forms of these materials are synthesized by both melt flux and solid-state routes, which provide some control over the crystallite size of the product. The melt flux process is time intensive, but it produces relatively large crystals at lower reaction temperatures. In comparison, the solid-state method is quicker yet requires higher reaction temperatures and yields smaller crystallites. Upon stirring in hot water, CaCuSi4O10 spontaneously exfoliates into monolayer nanosheets, which are characterized by TEM and PXRD. BaCuSi4O10 on the other hand requires ultrasonication in organic solvents to achieve exfoliation. Near infrared imaging illustrates that both the bulk and nanosheet forms of CaCuSi4O10 and BaCuSi4O10 are strong near infrared emitters. Aqueous CaCuSi4O10 and BaCuSi4O10 nanosheet dispersions are useful because they provide a new way to handle, characterize, and process these materials in colloidal form.

The described syntheses of CaCuSi4O10 and BaCuSi4O10 provide approximately 0.5 g of product per batch. Isolated yields of CaCuSi4O10 from the melt flux and solid-state syntheses typically range from 70-75% and 90-95%, respectively. For BaCuSi4O10, the isolated yields from the melt flux and solid-state syntheses typically range from 65-70% and 95-99%, respectively.
The textures of all of the prepared materials, as well as differences in the intensity of their blue color due to varying crystallite sizes, are visible by low magnification optical microscopy (Figures 3a-h). Scanning electron microscopy (SEM) images confirm that the solid-state method of synthesizing CaCuSi4O10 produces ~1-15 &#181;m primary crystallites (Figure 4b) whereas melt flux conditions lead to ~5-50 &#181;m crystallites (Figure 4a). Powder X-ray diffraction (PXRD) patterns for CaCuSi4O10 (Figures 5a and 5c) and BaCuSi4O10 (Figures 6a and 6c) showcase the composition and phase purity of these products.
Representative transmission electron microscopy (TEM) images show the nanosheet morphology of the exfoliated products (Figure 7). In addition, NIR photographic imaging shows the strong luminescence of both the bulk and exfoliated materials (Figure 8). A simple way to illustrate the solution processability of CaCuSi4O10 nanosheets is to prepare an aqueous ink (Figure 9) suitable for painting (Figure 10).
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/51686/51686fig1highres.jpg" src="/files/ftp_upload/51686/51686fig1.jpg" /> 
Figure 1.&#160;Photographs of the hand-ground starting materials.&#160;(a) CaCuSi4O10 melt flux, (b) CaCuSi4O10 solid-state, (c) BaCuSi4O10 melt flux, and (d) BaCuSi4O10 solid-state syntheses. <a href="https://www.jove.com/files/ftp_upload/51686/51686fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/51686/51686fig2highres.jpg" src="/files/ftp_upload/51686/51686fig2.jpg" /> 
Figure 2.&#160;Scanning Electron Microscopy. Images of the hand-ground starting materials for the (a) CaCuSi4O10 melt flux, (b) CaCuSi4O10 solid-state, (c) BaCuSi4O10 melt flux, and (d) BaCuSi4O10 solid-state syntheses. All samples were coated with gold prior to imaging. <a href="https://www.jove.com/files/ftp_upload/51686/51686fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" fo:content-width="5in" fo:src="/files/ftp_upload/51686/51686fig3highres.jpg" src="/files/ftp_upload/51686/51686fig3.jpg" /> 
Figure 3.&#160;Optical Microscopy. Bulk CaCuSi4O10 prepared by melt flux (a) and solid state (b) procedures. Bulk BaCuSi4O10 prepared by melt flux (c) and solid state (d) procedures. Exfoliated products (e-h) of (a-d), respectively. All images share the 1 mm scale bar show in panel (a). <a href="https://www.jove.com/files/ftp_upload/51686/51686fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" fo:content-width="5in" fo:src="/files/ftp_upload/51686/51686fig4highres.jpg" src="/files/ftp_upload/51686/51686fig4.jpg" /> 
Figure 4.&#160;Scanning Electron Microscopy. Images of bulk CaCuSi4O10 made by melt flux (a) and solid state (b) methods. Samples were coated with gold prior to imaging. <a href="https://www.jove.com/files/ftp_upload/51686/51686fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" fo:content-width="5in" src="/files/ftp_upload/51686/51686fig5highres.jpg" width="500" /> 
Figure 5.&#160;Powder X-Ray Diffraction: CaCuSi4O10. Patterns for bulk CaCuSi4O10 prepared by melt flux (a) and solid state (c) methods. Asterisks denote a silica impurity. Patterns for exfoliated CaCuSi4O10, (b) and (d), prepared from (a) and (c), respectively. <a href="https://www.jove.com/files/ftp_upload/51686/51686fig5highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" fo:content-width="5in" src="/files/ftp_upload/51686/51686fig6highres.jpg" width="500" /> 
Figure 6.&#160;Powder X-Ray Diffraction: BaCuSi4O10. Patterns for bulk BaCuSi4O10 prepared by melt flux (a) and solid state (c) methods. Asterisk denotes a silica impurity. Patterns for exfoliated BaCuSi4O10, (b) and (d), prepared from (a) and (c), respectively. <a href="https://www.jove.com/files/ftp_upload/51686/51686fig6highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" fo:content-width="5in" fo:src="/files/ftp_upload/51686/51686fig7highres.jpg" src="/files/ftp_upload/51686/51686fig7.jpg" /> 
Figure 7. Transmission Electron Microscopy. Representative images of exfoliated CaCuSi4O10 derived from bulk CaCuSi4O10 made by melt flux (a) or solid state (b) methods. Representative images of exfoliated BaCuSi4O10 derived from bulk BaCuSi4O10 made by melt flux (c) or solid state (d) methods. <a href="https://www.jove.com/files/ftp_upload/51686/51686fig7highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" fo:content-width="5in" fo:src="/files/ftp_upload/51686/51686fig8highres.jpg" src="/files/ftp_upload/51686/51686fig8.jpg" /> 
Figure 8.&#160;Near Infrared Imaging. Luminescence of bulk CaCuSi4O10 prepared by melt flux (a) and solid state (b) procedures. Luminescence of bulk BaCuSi4O10 prepared by melt flux (c) and solid state (d) procedures. Luminescence of the exfoliated products (e-h) of (a-d), respectively. Powder samples are contained within glass vials, and the entire set of samples was imaged at once. <a href="https://www.jove.com/files/ftp_upload/51686/51686fig8highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" fo:content-width="5in" fo:src="/files/ftp_upload/51686/51686fig9highres.jpg" src="/files/ftp_upload/51686/51686fig9.jpg" /> 
Figure 9.&#160;Photograph of a CaCuSi4O10 nanosheet ink in a vial.
<img alt="Figure 10" fo:content-width="5in" fo:src="/files/ftp_upload/51686/51686fig10highres.jpg" src="/files/ftp_upload/51686/51686fig10.jpg" /> 
Figure 10.&#160;Near Infrared Imaging. A rudimentary painting with the CaCuSi4O10 nanosheet ink that illustrates both its simple application and its luminescence properties.

Watch this Scientific Journal Video about Exfoliation of Egyptian Blue and Han Blue, Two Alkali Earth Copper Silicate-based Pigments at JoVE.com

Exfoliation of Egyptian Blue and Han Blue, Two Alkali Earth Copper Silicate-based Pigments

The preparation and exfoliation of CaCuSi4O10&#160;and BaCuSi4O10 are described. Upon stirring in hot water, CaCuSi4O10 spontaneously exfoliates into monolayers, whereas BaCuSi4O10&#160;requires ultrasonication in organic solvents. Near infrared (NIR) imaging illustrates the NIR emission properties of these materials, and aqueous dispersions of these nanomaterials are useful for solution processing.

In a visualized example of the ancient past connecting with modern times, we describe the preparation and exfoliation of CaCuSi4O10 and BaCuSi4O10, the ...

exfoliation-egyptian-blue-han-blue-two-alkali-earth-copper-silicate

Research

JoVE Journal

Chemistry

15.6K Views.  The University of Georgia. The preparation and exfoliation of CaCuSi4O10 and BaCuSi4O10 are described. Upon stirring in hot water, CaCuSi4O10 spontaneously exfoliates into monolayers, whereas BaCuSi4O10 requires ultrasonication in organic solvents. Near infrared (NIR) imaging illustrates the NIR emission properties of these materials, and aqueous dispersions of these nanomaterials are useful for soluti...

Video: Exfoliation of Egyptian Blue and Han Blue, Two Alkali Earth Copper Silicate-based Pigments

Quantifying Abdominal Pigmentation in Drosophila melanogaster

Pigmentation is a morphologically simple but highly variable trait that often has adaptive significance. It has served extensively as a model for understanding the development and evolution of morphological phenotypes. Abdominal pigmentation in Drosophila melanogaster has been particularly useful, allowing researchers to identify the loci that underlie inter- and intraspecific variations in morphology. Hitherto, however, D. melanogaster abdominal pigmentation has been largely assayed qualitatively, through scoring, rather than quantitatively, which limits the forms of statistical analysis that can be applied to pigmentation data. This work describes a new methodology that allows for the quantification of various aspects of the abdominal pigmentation pattern of adult D. melanogaster. The protocol includes specimen mounting, image capture, data extraction, and analysis. All the software used for image capture and analysis feature macros written for open-source image analysis. The advantage of this approach is the ability to precisely measure pigmentation traits using a methodology that is highly reproducible across different imaging systems. While the technique has been used to measure variation in the tergal pigmentation patterns of adult D. melanogaster, the methodology is flexible and broadly applicable to pigmentation patterns in myriad different organisms.

Quantifying Abdominal Pigmentation in Drosophila melanogaster

This work presents a method to quickly and precisely quantify the abdominal pigmentation of Drosophila melanogaster using digital image analysis. This method streamlines the procedures between phenotype acquisition and data analysis and includes specimen mounting, image acquisition, pixel value extraction, and trait measurement.

Pigmentation is a morphologically simple but highly variable trait that often has adaptive significance. It has served extensively as a model for ...

Genetics

Water in Oil Emulsions: A New System for Assembling Water-soluble Chlorophyll-binding Proteins with Hydrophobic Pigments

Chlorophylls (Chls) and bacteriochlorophylls (BChls) are the primary cofactors that carry out photosynthetic light harvesting and electron transport. Their functionality critically depends on their specific organization within large and elaborate multisubunit transmembrane protein complexes. In order to understand at the molecular level how these complexes facilitate solar energy conversion, it is essential to understand protein-pigment, and pigment-pigment interactions, and their effect on excited dynamics. One way of gaining such understanding is by constructing and studying complexes of Chls with simple water-soluble recombinant proteins. However, incorporating the lipophilic Chls and BChls into water-soluble proteins is difficult. Moreover, there is no general method, which could be used for assembly of water-soluble proteins with hydrophobic pigments. Here, we demonstrate a simple and high throughput system based on water-in-oil emulsions, which enables assembly of water-soluble proteins with hydrophobic Chls. The new method was validated by assembling recombinant versions of the water-soluble chlorophyll binding protein of Brassicaceae plants (WSCP) with Chl a. We demonstrate the successful assembly of Chl a using crude lysates of WSCP expressing E. coli cell, which may be used for developing a genetic screen system for novel water-soluble Chl-binding proteins, and for studies of Chl-protein interactions and assembly processes.

This manuscript describes a simple and high-throughput method for assembling water-soluble proteins with hydrophobic pigments that is based on water-in-oil emulsions. We demonstrate the effectiveness of the method in the assembly of native chlorophylls with four variants of recombinant water-soluble-chlorophyll binding proteins (WSCPs) of Brassica plants expressed in E. coli.

Chlorophylls (Chls) and bacteriochlorophylls (BChls) are the primary cofactors that carry out photosynthetic light harvesting and electron transport. ...

Probing the Limits of Egg Recognition Using Egg Rejection Experiments Along Phenotypic Gradients

Brood parasites lay their eggs in other females' nests, leaving the host parents to hatch and rear their young. Studying how brood parasites manipulate hosts into raising their young and how hosts detect parasitism provide important insights in the field of coevolutionary biology. Brood parasites, such as cuckoos and cowbirds, gain an evolutionary advantage because they do not have to pay the costs of rearing their own young. However, these costs select for host defenses against all developmental stages of parasites, including eggs, their young, and adults. Egg rejection experiments are the most common method used to study host defenses. During these experiments, a researcher places an experimental egg in a host nest and monitors how hosts respond. Color is often manipulated, and the expectation is that the likelihood of egg discrimination and the degree of dissimilarity between the host and experimental egg are positively related. This paper serves as a guide for conducting egg rejection experiments from describing methods for creating consistent egg colors to analyzing the findings of such experiments. Special attention is given to a new method involving uniquely colored eggs along color gradients that has the potential to explore color biases in host recognition. Without standardization, it is not possible to compare findings between studies in a meaningful way; a standard protocol within this field will allow for increasingly accurate and comparable results for further experiments.

This protocol provides guidelines for running egg rejection experiments: outlining techniques for painting experimental egg models to emulate the colors of natural bird eggs, conducting fieldwork, and analyzing the collected data. This protocol provides a uniform method for conducting comparable egg rejection experiments.

Brood parasites lay their eggs in other females' nests, leaving the host parents to hatch and rear their young. Studying how brood parasites manipulate ...

Behavior

Exfoliation and Analysis of Large-area, Air-Sensitive Two-Dimensional Materials

We describe methods for producing and analyzing large, thin flakes of air-sensitive two-dimensional materials. Thin flakes of layered or van der Waals crystals are produced using mechanical exfoliation, in which layers are peeled off a bulk crystal using adhesive tape. This method produces high-quality flakes, but they are often small and can be hard to find, particularly for materials with relatively high cleavage energies such as black phosphorus. By heating the substrate and the tape, two-dimensional material adhesion to the substrate is promoted, and the flake yield can be increased by up to a factor of ten. After exfoliation, it is necessary to image or otherwise analyze these flakes but some two-dimensional materials are sensitive to oxygen or water and will degrade when exposed air. We have designed and tested a hermetic transfer cell to temporarily maintain the inert environment of a glovebox so that air-sensitive flakes can be imaged and analyzed with minimal degradation. The compact design of the transfer cell is such that optical analysis of sensitive materials can be performed outside of a glovebox without specialized equipment or modifications to existing equipment.

A method for exfoliating large thin flakes of air sensitive two-dimensional materials and safely transporting them for analysis outside of a glovebox is presented.

We describe methods for producing and analyzing large, thin flakes of air-sensitive two-dimensional materials. Thin flakes of layered or van der Waals ...

Electrochemical Exfoliation to Produce High-Quality Black Phosphorus

To obtain high-quality two-dimensional (2D) materials from the bulky crystals, delamination under an externally controlled stimulus is crucial. Electrochemical exfoliation of layered materials requires simple instrumentation yet offers high-quality exfoliated 2D materials with high yields and features straightforward upscalability; therefore, it represents a key technology for advancing fundamental studies and industrial applications. Moreover, the solution processability of functionalized 2D materials enables the fabrication of (opto)electronic and energy devices via different printing technologies such as inkjet printing and 3D printing. This paper presents the electrochemical exfoliation protocol for the synthesis of black phosphorus (BP), one of the most promising emerging 2D materials, from its bulk crystals in a step-by-step manner, namely, cathodic electrochemical exfoliation of BP in the presence of N(C4H9)4&#8729;HSO4 in propylene carbonate, dispersion preparation by sonication and subsequent centrifugation for the separation of flakes, and morphological characterization by scanning electron microscopy (SEM), atomic force microscopy (AFM), and transmission electron microscopy (TEM).

We present a step-by-step procedure for electrochemical exfoliation of black phosphorus (BP), one of the most promising emerging 2D materials with applications in (opto)electronics, from its bulk crystals, as well as the morphological characterization by scanning electron microscopy, atomic force microscopy, and transmission electron microscopy.

To obtain high-quality two-dimensional (2D) materials from the bulky crystals, delamination under an externally controlled stimulus is crucial. ...

In Vitro Reconstitution of Light-harvesting Complexes of Plants and Green Algae

In plants and green algae, light is captured by the light-harvesting complexes (LHCs), a family of integral membrane proteins that coordinate chlorophylls and carotenoids. In vivo, these proteins are folded with pigments to form complexes which are inserted in the thylakoid membrane of the chloroplast. The high similarity in the chemical and physical properties of the members of the family, together with the fact that they can easily lose pigments during isolation, makes their purification in a native state challenging. An alternative approach to obtain homogeneous preparations of LHCs was developed by Plumley and Schmidt in 19871, who showed that it was possible to reconstitute these complexes in vitro starting from purified pigments and unfolded apoproteins, resulting in complexes with properties very similar to that of native complexes. This opened the way to the use of bacterial expressed recombinant proteins for in vitro reconstitution. The reconstitution method is powerful for various reasons: (1) pure preparations of individual complexes can be obtained, (2) pigment composition can be controlled to assess their contribution to structure and function, (3) recombinant proteins can be mutated to study the functional role of the individual residues (e.g., pigment binding sites) or protein domain (e.g., protein-protein interaction, folding). This method has been optimized in several laboratories and applied to most of the light-harvesting complexes. The protocol described here details the method of reconstituting light-harvesting complexes in vitro currently used in our laboratory, and examples describing applications of the method are provided.

In Vitro Reconstitution of Light-harvesting Complexes of Plants and Green Algae

This protocol details the reconstitution of light-harvesting complexes in vitro. These integral membrane proteins coordinate chlorophylls and carotenoids and are responsible for harvesting light in higher plants and green algae.

In plants and green algae, light is captured by the light-harvesting complexes (LHCs), a family of integral membrane proteins that coordinate chlorophylls ...

Biology

A Method for Extracting Pigments from Squid Doryteuthis pealeii

Cephalopods can undergo rapid and adaptive changes in dermal coloration for sensing, communication, defense, and reproduction purposes. These capabilities are supported in part by the areal expansion and retraction of pigmented organs known as chromatophores. While it is known that the chromatophores contain a tethered network of pigmented granules, their structure-function properties have not been fully detailed. We describe a method for isolating the nanostructured granules in squid Doryteuthis pealeii chromatophores and demonstrate how their associated pigments can be extracted in acidic solvents. To accomplish this, the chromatophore containing dermal layer is first manually isolated using a superficial dissection, and the pigment granules are removed using sonication, centrifugation, and washing cycles. Pigments confined within the purified granules are then extracted via acidic methanol solutions, leaving nanostructures with smaller diameters that are void of visible color. This extraction procedure produces a 58% yield of soluble pigments isolated from granules. Using this method, the composition of the chromatophore pigments can be determined and used to provide insight into the mechanism of adaptive coloration in cephalopods.

A Method for Extracting Pigments from Squid Doryteuthis pealeii

A protocol for the extraction of pigments from the nanostructured granules in squid Doryteuthis pealeii chromatophores is presented.

Cephalopods can undergo rapid and adaptive changes in dermal coloration for sensing, communication, defense, and reproduction purposes. These capabilities ...

Column Chromatography

Source: Lara Al Hariri and Ahmed Basabrain at the University of Massachusetts Amherst, MA, USA
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse="">Column Chromatography of Food Dye<button class="expand">Expand</button>
	In this experiment, you&#39;ll use column chromatography to separate green food coloring into its component blue and yellow dyes, called erioglaucine and tartrazine. First, you&#39;ll use 95% ethanol to elute the dye with a weaker affinity for silica gel. Then, you&#39;ll flush the second dye from the column with water. To recover the purified dyes, you will collect each band of dye as it leaves the column. These collected volumes are called fractions.
	<ul class="lab-items">
		<li data-sub-note-item="1-2">
		<div class="lab-step">To begin, put on a lab coat, safety glasses, and a pair of nitrile gloves.</div>
		</li>
		<li data-sub-note-item="1-3">
		<div class="lab-step">Bring a column or burette to the central supply area. Obtain a piece of cotton or glass wool and press it into the top of your column. Use a long, blunt-ended wire or rod to push it down to plug the column just above the stopcock. Note: The cotton must be stable, but don&#39;t pack it too densely, because the solvent needs to pass through it easily. Make sure that the stopcock can turn freely without catching on the plug.</div>
		</li>
		<li data-sub-note-item="1-4">
		<div class="lab-step">Attach a burette clamp to a lab stand and clamp the column upright with about 4 in of space underneath it. Make sure that the stopcock is closed.</div>
		</li>
		<li data-sub-note-item="1-5">
		<div class="lab-step">Label a 100-mL beaker as &#8216;waste&#8217; and place it under the column.</div>
		</li>
		<li data-sub-note-item="1-6">
		<div class="lab-step">Next, measure 1 g of sand in a tared weighing boat and bring it back to the bench.</div>
		</li>
		<li data-sub-note-item="1-7">
		<div class="lab-step">Then, mark the column about 0.5 cm above the top of the cotton plug.</div>
		</li>
		<li data-sub-note-item="1-8">
		<div class="lab-step">Place a small funnel in the top of the column and pour enough sand through the funnel so that there is a layer 0.5 cm deep over the plug. This will keep the silica gel from passing through the plug into your fractions.</div>
		</li>
		<li data-sub-note-item="1-9">
		<div class="lab-step">Remove the funnel and gently tap the column to evenly distribute the sand.</div>
		</li>
		<li data-sub-note-item="1-10">
		<div class="lab-step">Now, prepare your eluents. Label a 50-mL beaker and a 50-mL graduated cylinder as &#8216;water&#8217;.</div>
		</li>
		<li data-sub-note-item="1-11">
		<div class="lab-step">Measure 50 mL of deionized water with the graduated cylinder and pour it into the beaker.</div>
		</li>
		<li data-sub-note-item="1-12">
		<div class="lab-step">Then, label another 50-mL beaker and a 50-mL graduated cylinder as &#8216;ethanol&#8217;. Measure 50 mL of 95% ethanol and pour it into this beaker.</div>
		</li>
		<li data-sub-note-item="1-13">
		<div class="lab-step">Now, obtain 4 or 5 clean Pasteur pipettes and a pipette bulb. Gently pipette 7 &#8211; 8 mL of ethanol along the inner walls of the column. This layer of solvent will let you pour this silica gel slurry into the column without disturbing the sand.</div>
		</li>
		<li data-sub-note-item="1-14">
		<div class="lab-step">Open the stopcock and drain ~1 mL of ethanol into the waste beaker to flush air from the stopcock and to confirm that the solvent flows easily through the plug. Make sure that there are no dry patches in the sand and the plug and gently tap the side of the column to remove trapped air bubbles.</div>
		</li>
		<li data-sub-note-item="1-15">
		<div class="lab-step">Next, place a clean funnel in the top of the column and label a third 50-mL beaker &#8216;silica gel slurry in ethanol&#8217;.</div>
		</li>
		<li data-sub-note-item="1-16">
		<div class="lab-step">Carefully weigh 5 g of silica gel powder and pour it into your labeled beaker.</div>
		</li>
		<li data-sub-note-item="1-17">
		<div class="lab-step">Then, bring the silica gel, a glass rod, and your ethanol graduated cylinder to the ethanol dispensing hood.</div>
		</li>
		<li data-sub-note-item="1-18">
		<div class="lab-step">Measure 18 mL of 95% ethanol and pour it into the beaker. Stir the mixture with the glass rod until the texture is consistent and no dry patches or clumps of powder remain. This usually takes ~1 min. Note: The slurry should be thick, like batter, but it should still flow easily. If the slurry is too stiff, it could clog the funnel or trap air bubbles in the column.</div>
		</li>
		<li data-sub-note-item="1-19">
		<div class="lab-step">Once the powder is completely suspended in ethanol, bring it to your bench. Stir the slurry until its texture is consistent again and immediately pour it into the column.</div>
		</li>
		<li data-sub-note-item="1-20">
		<div class="lab-step">Gently tap the sides of the column as the silica gel settles to get rid of air bubbles.</div>
		</li>
		<li data-sub-note-item="1-21">
		<div class="lab-step">Rinse the beaker with 1 or 2 mL of ethanol, and pour the rinse into the column. Then, remove the funnel and set it aside.</div>
		</li>
		<li data-sub-note-item="1-22">
		<div class="lab-step">Pipette ethanol along the inside of the column to wash down particles stuck to the walls. Then, drain the solvent until the meniscus is just barely above the top of the silica gel.</div>
		</li>
		<li data-sub-note-item="1-23">
		<div class="lab-step">Now, use a 5- or 10-mL graduated cylinder to obtain about 0.8 mL of the green food dye solution.</div>
		</li>
		<li data-sub-note-item="1-24">
		<div class="lab-step">Use a clean pipette to gently apply 5 to 7 drops of the dye solution evenly across the top of the silica gel. If necessary, add a few drops of ethanol to keep the silica gel covered.</div>
		</li>
		<li data-sub-note-item="1-25">
		<div class="lab-step">While you wait for the dye to soak in, label two clean 50-mL beakers as &#8216;ethanol fraction&#8216; and &#8216;water fraction&#8217;. Exchange the waste beaker under the column with the ethanol fraction beaker.</div>
		</li>
		<li data-sub-note-item="1-26">
		<div class="lab-step">Next, fill the column with ethanol, pipetting the first mL along the inner walls to avoid disturbing the silica gel.</div>
		</li>
		<li data-sub-note-item="1-27">
		<div class="lab-step">Then, open the stopcock partway and adjust it until the solvent level decreases ~1 mL every 15 &#8211; 20 s.</div>
		</li>
		<li data-sub-note-item="1-28">
		<div class="lab-step">Periodically refill the column to keep the ethanol flowing. Make sure that there is at least 1 cm of solvent above the silica gel at all times.</div>
		</li>
		<li data-sub-note-item="1-29">
		<div class="lab-step">Monitor the yellow and blue bands as they separate. Note: If you need to step away, close the stopcock so that the column won&#39;t run dry while you&#39;re gone. Avoid stopping for more than 1 min at a time because the compound bands will start broadening. It usually takes about 20 &#8211; 35 min for the faster-moving dye to leave the column.</div>
		</li>
		<li data-sub-note-item="1-30">
		<div class="lab-step">Once you have collected all of the faster-moving dye in the ethanol fraction beaker, close the stopcock, set the ethanol fraction aside, and put the waste beaker under the column.</div>
		</li>
		<li data-sub-note-item="1-31">
		<div class="lab-step">Drain the solvent until the meniscus is just above the top of the silica gel. Then, exchange the waste beaker for the water fraction beaker.</div>
		</li>
		<li data-sub-note-item="1-32">
		<div class="lab-step">Use a clean pipette to fill the top of the column with deionized water. Then, open the stopcock just enough to let water flow through the column at a rate of about 1 mL every 15 &#8211; 20 s.</div>
		</li>
		<li data-sub-note-item="1-33">
		<div class="lab-step">Refill the column with water as needed until the second band of color has completely left the column. This usually takes another 15 &#8211; 25 min. Then, close the stopcock and exchange the water fraction beaker for the waste beaker.</div>
		</li>
		<li data-sub-note-item="1-34">
		<div class="lab-step">Now, drain the column completely and leave the stopcock open.</div>
		</li>
		<li data-sub-note-item="1-35">
		<div class="lab-step">Record the colors of the ethanol and water fractions in your lab notebook.
		<h4>Table 1: Column Chromatography of Food Dye</h4>
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
			<tbody>
				<tr>
					<td></td>
					<td>Color </td>
					<td>Volume (mL) </td>
				</tr>
				<tr>
					<td>Ethanol fraction</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td>Water fraction</td>
					<td></td>
					<td></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www.jove.com/CDNSource/lm/labs/63/Table_1._Column_chromatography_of_food_dye.xlsx" target="_blank">Click Here to download Table 1</a></div>
		</li>
		<li data-sub-note-item="1-36">
		<div class="lab-step">Pour the fractions into the corresponding 50 mL graduated cylinders and record the volume of each fraction. Then, close the stopcock and bring the column to a fume hood.</div>
		</li>
		<li data-sub-note-item="1-37">
		<div class="lab-step">Clamp the column upright in the hood so that the silica gel can dry. Your instructor will either tell you how to clean the column or take care of it after the lab.</div>
		</li>
		<li data-sub-note-item="1-38">
		<div class="lab-step">Now, flush the collected fractions and leftover ethanol, water, and dye solution down the drain and clean the rest of your glassware as usual.</div>
		</li>
		<li data-sub-note-item="1-39">
		<div class="lab-step">Discard used Pasteur pipettes in the glass waste. Throw out used paper towels and weighing boat in the lab trash and clean the benchtop before you leave.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="2" data-parent-collapse="">Results<button class="expand">Expand</button>
	In this experiment, you saw that ethanol eluted the blue erioglaucine in a timely manner, but it wasn&#39;t an effective eluent for the yellow tartrazine. Paper chromatography of the green dye shows similar results. Thus, we conclude that erioglaucine has a lower affinity for silica gel than tartrazine does.
	Now, consider the advantages of using two solvents to elute the dye separately. Water is more polar than ethanol, so we assume that it would elute both dyes faster than ethanol alone would. While you also would have used less solvent overall if you had only used water, the separation would have been worse, making it harder to collect pure fractions.
	Alternatively, you could have kept using ethanol after you collected the erioglaucine, but it would have taken a long time and a lot of ethanol to elute tartrazine that way. Switching the eluent to water after eluting erioglaucine saved time and minimized solvent waste without sacrificing purity.
	<ul class="lab-items">
	</ul>
	</li>
</ol>

Column Chromatography: Experimental Setup and Separation - Procedure

Chromatografia kolumnowa

Source: Lara Al Hariri and Ahmed Basabrain at the University of Massachusetts Amherst, MA, USA

	Column Chromatography of Food DyeExpand
	In this ...

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JoVE Encyclopedia of Experiments

Cancer Research

Urinary Tract Cancer

Ex vivo studies

Photosynthesis

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Plant Pigment Chromatography
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">NOTE: In this experiment you will separate pigments from spinach leaves using chromatography paper. Individual pigments travel along the paper at different rates and may have different colors. By calculating the relative distance the pigments travel, their resolution factor, and comparing them with literature values, you can identify different pigments. HYPOTHESES: In this exercise the experimental hypothesis is that there will be multiple pigments within the spinach leaves that absorb different wavelengths of sunlight. The null hypothesis is that there is only one type of pigment within the spinach leaf.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Use a pencil to make a line two centimeters from one end of the chromatography paper.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Then, lay a pipe cleaner horizontally across the top of a clean 400 mL beaker.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Place the pencil-marked side of the chromatography strip at the bottom of the beaker.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Next, wrap the paper around the pipe cleaner so that the bottom edge is barely touching the bottom of the beaker and then secure it with a paperclip.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">When the paper is secured around the pipe cleaner, remove it from the beaker, and then place a patted-dry spinach leaf over the marked line on the chromatography paper.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Roll a coin over the spinach leaf along the pencil line going back and forth multiple times and applying steady pressure. When the leaf is removed, a green line should be clearly present.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Next, place 8 mL of chromatography solvent in the beaker.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Lower the chromatography strip into the beaker so that the edge of the paper touches the solvent but the green line does not. Adjust the pipe cleaner if needed.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">Without disturbing the beaker, observe the solvent as it moves up the paper and the individual pigments separate.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">When the solvent has traveled half way up the chromatography paper, which will take approximately 10 minutes, and the pigments have separated into well-defined bands, remove the paper from the beaker.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Mark how far the solvent traveled with a pencil and then allow the paper to dry. NOTE: The solvent evaporates quickly.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">Next, record the number of visible bands and describe their color and relative size.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Measure how far the solvent and pigments traveled, and record this information for each pigment in Table 1. <a href="https://www.jove.com/CDNSource/lm/labs/7/Table_1_Photosynthesis.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Dispose of the chromatography solvent in a waste container under a fume hood. Throw the chromatography strips into the regular trash, and then clean the beakers with soap and water.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Floating Leaf Discs in a Vacuum
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">NOTE: In this experiment you will indirectly observe photosynthesis and cellular respiration using a floating leaf disc in a solution. During photosynthesis, air bubbles will cause the leaves to float, and during respiration, the discs will sink. HYPOTHESES: In this exercise, the experimental hypothesis is that the leaf discs will have a greater rate of photosynthesis in the bicarbonate solution, because bicarbonate provides added CO2 to fuel photosynthesis, causing more leaf discs to float. Additionally, all of the discs will sink in dark conditions as they perform cellular respiration. The null hypothesis is that there will be no difference in the rate of photosynthesis, and therefore the number of floating discs, between the bicarbonate and water, or light and dark treatments.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">To place leaf discs under vacuum, first remove the plungers from two 20 mL syringes, and then place 10 leaf discs inside each syringe tube. Label one syringe &#8220;bicarbonate&#8221;, and label the other syringe &#8220;water&#8221;.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Replace the plungers and push the plunger until only a small amount of air remains in the syringe. Take care not to damage the leaf discs.
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Pull 5 mL of the bicarbonate solution into one of the syringes. Invert and swirl the syringe to suspend the leaf discs in solution.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">Push as much air out as possible without expelling the solution or damaging the leaf discs.
</div></li><li data-sub-note-item="2-6"><div class="lab-step">Then pull 5 mL of the water solution into the other syringe and swirl it as previously described (step 3).
</div></li><li data-sub-note-item="2-7"><div class="lab-step">To create a vacuum, hold one finger over the tip of the syringe while pulling back on the plunger. Hold this for 10 seconds while swirling the syringe to keep the leaf discs in suspension.
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Then, release the vacuum. NOTE: The discs should have absorbed the solution into the air spaces in their tissues and you should see them sink. If the discs don&#39;t sink, you can repeat the vacuum creation up to three times.
</div></li><li data-sub-note-item="2-9"><div class="lab-step">Next, add 50 mL of bicarbonate solution to a plastic cup or a glass beaker, and then gently add the discs from the bicarbonate vacuum syringe.
</div></li><li data-sub-note-item="2-10"><div class="lab-step">For the control, add the same amount of water to an identical cup, and then add the leaf discs from the water vacuum syringe. Label the containers appropriately.
</div></li><li data-sub-note-item="2-11"><div class="lab-step">Place both cups under a light source.
</div></li><li data-sub-note-item="2-12"><div class="lab-step">Every five minutes record the number of discs floating on the surface of the cup in Table 3 until 20 minutes have passed. <a href="https://www.jove.com/CDNSource/lm/labs/7/Table_3_Photosynthesis.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 3</a>
</div></li><li data-sub-note-item="2-13"><div class="lab-step">Next, remove the cups from the light source and then swirl them so that the discs at the surface intermix with any gases also at the surface.
</div></li><li data-sub-note-item="2-14"><div class="lab-step">Move the cups to a dark place. Every five minutes record the number of leaf discs floating at the surface until 20 minutes have passed. Swirl the cup each time before placing it back in the dark.
</div></li><li data-sub-note-item="2-15"><div class="lab-step">To clean up, dispose of the leaf discs in the trash, and pour the bicarbonate solution down the drain. Wash the syringes and cups thoroughly.
</div></li></ul></li><li data-note-item="3" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="3-1"><div class="lab-step">NOTE: In the first experiment, you observed how far pigments from spinach leaves traveled on chromatography paper. Different pigments absorb light at different wavelengths.
</div></li><li data-sub-note-item="3-2"><div class="lab-step">Using colored pens or pencils, draw the positions of the pigment bands and the solvent on Figure 3.
</div></li><li data-sub-note-item="3-3"><div class="lab-step">Calculate the retention factor, or Rf values for the pigments, which is done by dividing the distance the pigment in question moved up the paper from the line by the distance the solvent moved up the paper from the line.
</div></li><li data-sub-note-item="3-4"><div class="lab-step">Compare your calculated Rf values to those in Table 2 to determine the identity of the pigment. <a href="https://www.jove.com/CDNSource/lm/labs/7/Table_2_Photosynthesis.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 2</a>
</div></li><li data-sub-note-item="3-5"><div class="lab-step">Record these data in Table 1. NOTE: In the second experiment, you observed floating and sinking leaf discs as an indirect measurement of photosynthesis and respiration.
</div></li><li data-sub-note-item="3-6"><div class="lab-step">Graph the results with time and minutes on the x-axis and number of floating discs on the y-axis. Use two different lines to represent the water control and the bicarbonate treatment.
</div></li><li data-sub-note-item="3-7"><div class="lab-step">Add a line to the graph to indicate the point where the discs were removed from the light condition and placed into the dark.
</div></li><li data-sub-note-item="3-8"><div class="lab-step">Next, starting with the bicarbonate condition, use the graph to determine the point at which 50% of the leaf discs were floating. This is referred to as the effective time, or ET50. NOTE: You will notice that the discs likely hit the 50% floating mark once in the light condition and then again in the dark condition.
</div></li><li data-sub-note-item="3-9"><div class="lab-step">Your water samples may or may not have reached the ET50 mark. If they did, add the line for this sample also.
</div></li><li data-sub-note-item="3-10"><div class="lab-step">Finally, compare your ET50 values and graphs with the rest of the class.
</div></li></ul></li></ol>

Studying Photosynthesis by Leaf Disc Experiment-Procedure

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Plant Pigment Chromatography
ExpandNOTE: In this experiment you will separate pigments from spinach leaves using chromatography paper. Individual pigments ...

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