The Research Center Pharmaceutical Engineering (RCPE) is funded within the framework of COMET - Competence Centers for Excellent Technologies by BMK, BMDW, Land Steiermark and SFG. The COMET program is managed by the FFG.

1. Differential scanning calorimetry (DSC)
<ol>
	<li>Instrument preparation
	<ol>
		<li>Use a differential scanning calorimeter equipped with an intracooler, an autosampler, and the software for instrument control and data analysis.</li>
		<li>Switch on the nitrogen gas supply and set the pressure between 0.2&#8211;0.5 bar and power up the DSC instrument and the automatic sample changer.</li>
		<li>Open the software and activate the standby mode by clicking on the Yes button. Allow equilibration of the device for at least one hour</li>
		<li>Purge the furnace with nitrogen, click on the New Method icon and go to Method Definition. Activate the Temperature Modulation option in the overview window.&#160; Go to header tab and select the method by clicking on &#8220;sample&#8221;.</li>
		<li>Go to the tab Temperature Program, select Purge 2 MFC and on Protective MFC, both at 50 mL/min.</li>
		<li>Insert the following measurement method: Standby at 20 &#176;C, heating cycle at 5 K/min from 20 &#176;C to above the melting temperature of lipid, isothermal holding at this temperature for 5 min, cooling cycle to 0 &#176;C at 5 K/min to -20 &#176;C, final emergency reset temperature at a temperature 10 degrees &#176;C above the highest temperature of the program, and final standby temperature at 20 &#176;C.</li>
		<li>Go to the calibration tab and select the appropriate temperature and sensitivity file. Save the method</li>
	</ol>
	</li>
	<li>Sample preparation and measurement
	<ol>
		<li>Weigh 3&#8211;4 mg of each sample into aluminum crucibles. Record the exact weight loaded into each crucible and seal the aluminum crucible with a pierced lid.</li>
		<li>Place the crucibles in the autosampler tray and activate the autosampler mode in the software and load related method for each sample.</li>
		<li>Fill out the sample position, sample name, and weight of each sample, and the position of reference crucible in the Sample Tray View window and start the measurements.</li>
	</ol>
	</li>
	<li>Data analysis
	<ol>
		<li>Open the raw data using the software for data analysis and plot the temperature versus heat flow, by clicking on the button &#8220;X-time / X-temperature&#8221;</li>
		<li>On the popped out window, click on &#8220;Hide isothermal segments&#8221;. On the left side of the screen, select only the curves that are to be analyzed (e.g. unclick the &#8220;Additional&#8221; data).</li>
		<li>Check the thermal behavior of lipids as endothermic and exothermic events of absorbed or released energy in the form of heat, respectively, as a function of temperature.</li>
		<li>Click on the curve, followed by Evaluation and Area, to calculate the enthalpy of fusion as the area under the curve of endotherms.</li>
		<li>Select the integration boundaries by moving the vertical lines around 2 to 3 degrees Celsius before and after the onset and endpoint of the peak.</li>
		<li>Select a linear baseline for the peak integration. The area between the curve and the baseline is proportional to the change in enthalpy. Click on Apply to finish the calculation. Similarly, calculate the enthalpy of crystallization as the area under the curve of exotherms</li>
		<li>Determine the onset of melting temperature (To) by clicking on the curve to be analyzed and then on Evaluation and Onset.</li>
		<li>Select the quantification boundaries by moving the vertical lines to the most straight section of the curve. This is usually around 5-10&#176;C before and after the peak. Then, determine the melting temperature by clicking on the curve to be analyzed, followed by Evaluation and Peak. The obtained value is the peak maximum.</li>
	</ol>
	</li>
</ol>
2. Small- and wide-angle x-ray scattering (SWAXS)
<ol>
	<li>Instrument preparation
	<ol>
		<li>Use an x-ray scattering system, composing a point-focusing camera fixed to a sealed-tube x-ray generator and equipped with a control unit and related software.</li>
		<li>Use cooper (&#955; = 1.54 &#197;) at 50 kV and 1 mA as an x-ray source and two linearly positioned sensitive detectors to cover both small- and wide-angle x-ray scattering regions.</li>
		<li>Ensure safety requirements to protect from x-ray exposure.</li>
		<li>&#160;Switch on the cooling water system on the control unit, the vacuum pump, the gas valves, and the power and safety control system.</li>
		<li>Switch on the voltage control and purging valves for the detectors at a gas flow between 10&#8211;20 mL/min.</li>
		<li>Switch on the x-ray tube and the standby option and wait approximately 10 min. Switch off the standby mode and power up the x-ray tube to full power (&#62;50kV) and wait at least 30 minutes.</li>
		<li>Start the control software and click on RESET TPF. Choose Tugsten filter and set position. Go to Position to fix the position of the Tungsten filter</li>
	</ol>
	</li>
	<li>Sample preparation and measurement
	<ol>
		<li>Make sure that the samples are available as fine powder. If necessary, grind the samples gently at low temperatures to provide a fine powder.</li>
		<li>Fill the samples into special glass capillaries with an external diameter of approximately 2 mm, avoiding any air entrapment in the capillaries. Seal the glass capillary with wax and place it carefully into the capillary holder.</li>
		<li>Switch on the motor for sample rotation and close the vacuum valve until the pressure is beneath 5 mbar.</li>
		<li>In the software, fix the measurements setting by selecting a position resolution of 1024. Fix the exposure time to 1200 s.</li>
		<li>Setup the energy limits by clicking on the tap Tools, then click on energy and resolution, and click on restart. Setup the energy limits to a suitable range between 400-900.</li>
		<li>Open the safety shutter and start the measurements. Ensure that the measurement window shows a maximum of 80 counts per second. If this is not given, adapt the filter position.</li>
	</ol>
	</li>
	<li>Data analysis
	<ol>
		<li>Export the data as p00 files. The data consists of the intensity of transmission and absorption versus the channel number and the diffraction angle.</li>
		<li>Transfer the data for evaluation to a statistical software and correct the data by normalizing the intensities using the scattering mass measured with the Tungsten filter.</li>
		<li>Create a plot of normalized intensity versus two times the diffraction angle [(2&#920;) 2xtheta].</li>
		<li>Use the function of &#8220;screen reader&#8221; to find diffraction peaks in SAXS and WAXS regions.</li>
		<li>Apply Bragg&#39;s equation to compute the diffraction peaks with maximum intensity into short and long d-spacing for WAXS and SAXS regions, respectively.</li>
		<li>Calculate the ratios of the peak position of the SAXS region to find out the crystal symmetry of the lipids (e.g., lamellar, hexagonal, cubic).</li>
		<li>Use the main diffraction peak of the SAXS region to quantify the crystallite thickness (D). Fit the peak into a Gaussian function via classical least squares and obtain the FWHM by clicking on Analysis, Peaks and baselines, Peak analyzer, Open dialog.</li>
		<li>On the popped out window, select the option &#8220;Fit Peaks Pro&#8221;. Select a constant baseline with y = 0, and select the main diffraction peak of the SAXS region and click on &#8220;Fit control&#8221; to select the peak fit parameters.</li>
		<li>Choose the GaussAmp function. Set the parameters y_0, xc_1 and A_1 as fixed and obtain the FWHM from the fit. Use the Scherrer equation to compute the crystallite thickness.</li>
	</ol>
	</li>
</ol>
3. Dissolution tests
<ol>
	<li>API release from coated multiparticulate systems
	<ol>
		<li>Use USP apparatus 2 (paddle) for dissolution studies.</li>
		<li>Fill the dissolution test vessels with phosphate buffer pH 6.8, and heat to 37 &#176;C.</li>
		<li>Weigh triplicates of samples of coated particles equivalent to a single dose of API, and place the samples into dissolution test vessels.</li>
		<li>Start the paddle at a speed of 100 rpm.</li>
		<li>Set the auto-sampler to take samples of 1 mL at the following sampling points: 30 min, 60 min, 90 min, 2 h, 4 h, 6 h, 8 h, 10 h, 12 h, 18 h, and 24 h.</li>
		<li>Analyze the samples via a suitable HPLC method5,7,17.</li>
		<li>Analyze the data by plotting the cumulative API release versus time.</li>
		<li>Carry out the experiments for stored samples under long-term (25 &#176;C, 60% relative humidity) and accelerated (40 &#176;C and 70% relative humidity).</li>
	</ol>
	</li>
	<li>API release from solid lipid-nanoparticles (SLN)
	<ol>
		<li>Prepare simulated lung fluid (SLF) by mixing 0.02% (w/w) of dipalmitoylphosphatidylcholine in Dulbecco&#39;s phosphate buffer saline (D-PBS), with the following composition: KCl (2.683 mM), KH2PO4 (1.47 mM), NaCl (136.893 mM), Na2HPO4&#183;2H2O (8.058 mM), CaCl2&#183;2H2O (0.884 mM), and MgCl2&#183;6H2O (0.492 mM). Pre-heat it at 37 &#176;C.</li>
		<li>Use dialysis cassettes with a cellulose membrane bag of a controlled cut-off of 7,000 Da in triplicate for each sample.</li>
		<li>Assign one dialysis cassette for each sampling time: 0.5 h, 1.5 h, 3 h, 5 h, 7 h, and 24 h. Hydrate the dialysis cassettes for 2 min by immersing them in SLF. Then, carefully dry their surface with soft paper towels.</li>
		<li>Inject 3 mL of the sample (lipid-nanosuspension), equivalent to 600 &#181;g of dexamethasone, into each cassette.</li>
		<li>Immerse each cassette in 200 mL of SLF at 37 &#176;C (sink conditions) and agitate the system at 125 rpm.</li>
		<li>Take 200 mg samples from inside the cassette using a syringe at each determined sampling time.</li>
		<li>Determine the API content using the developed HPLC-MS method18.</li>
		<li>Calculate the API released from SLN by mass balance, according to18, briefly as the difference between the total amount of API in SLN and the remaining amount of API after sampling.</li>
		<li>Repeat the process for stored samples.</li>
	</ol>
	</li>
</ol>

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L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thermal+and+structural+behaviour+of+crude+palm+oil:+Crystallisation+at+very+low+cooling+rate.">Thermal and structural behaviour of crude palm oil: Crystallisation at very low cooling rate.</a> European Journal of Lipid Science and Technology. 109 (4), 410-421 (2007).</li><li>Askin, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simultaneous+differential+scanning+calorimetry-X-ray+diffraction+study+of+olanzapine+crystallization+from+amorphous+solid+dispersions.">A simultaneous differential scanning calorimetry-X-ray diffraction study of olanzapine crystallization from amorphous solid dispersions.</a> Molecular Pharmaceutics. 17 (11), 4364-4374 (2020).</li><li>Clout, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+differential+scanning+calorimetry+-+synchrotron+X-ray+powder+diffraction:+A+powerful+technique+for+physical+form+characterization+in+pharmaceutical+materials.">Simultaneous differential scanning calorimetry - synchrotron X-ray powder diffraction: A powerful technique for physical form characterization in pharmaceutical materials.</a> Analytical Chemistry. 88 (20), 10111-10117 (2016).</li><li>Jendrzejewska, I., Goryczka, T., Pietrasik, E., Klimontko, J., Jampilek, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=X-ray+and+thermal+analysis+of+selected+drugs+containing+acetaminophen.">X-ray and thermal analysis of selected drugs containing acetaminophen.</a> Molecules. 25 (24), 5909 (2020).</li><li>Righetti, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crystallization+of+Polymers+Investigated+by+Temperature-Modulated+DSC.">Crystallization of Polymers Investigated by Temperature-Modulated DSC.</a> Materials. 10 (4), 442 (2017).</li><li>Sauer, B. B., Kampert, W. G., Neal Blanchard, E., Threefoot, S. A., Hsiao, B. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temperature+modulated+DSC+studies+of+melting+and+crystallization+in+polymers+exhibiting+multiple+endotherms.">Temperature modulated DSC studies of melting and crystallization in polymers exhibiting multiple endotherms.</a> Polymer. 41 (3), 1099-1108 (2000).</li><li>Ali, F., Kumar, R., Lal Sahu, P., Singh, G. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physicochemical+characterization+and+compatibility+study+of+roflumilast+with+various+pharmaceutical+excipients.">Physicochemical characterization and compatibility study of roflumilast with various pharmaceutical excipients.</a> Journal of Thermal Analysis and Calorimetry. 130, 1627-1641 (2017).</li></ol>

Authors disclose any and all conflicts of interest.

Powder x-ray diffraction and DSC were described in this manuscript as gold standards for the solid-state analysis of LBEs. Powder x-ray diffraction has the outstanding advantage of processing the measurements in situ, with minimum solid-state manipulation of samples during the measurements. Moreover, the same-filled capillaries can be stored under different conditions after initial measurements to investigate the solid-state alteration during storage. In this work, we focused on the wide- and small-angle regions...

Lipids are a class of materials that contain long-chain aliphatic hydrocarbons and their derivatives. They cover a broad range of chemical structures, including fatty acids, acylglycerols, sterols and sterol esters, waxes, phospholipids, and sphingolipids1. The use of lipids as pharmaceutical excipients started in 1960 for embedding drugs in a wax matrix to provide sustained release formulations2. Since then, lipid-based excipients (LBEs) have gained extensive attention for diverse applications, such as modified drug release, taste-masking, drug encapsulation, and enhanced drug bioavailability. LBEs can be applied in a broad range of pharmaceutical dosage forms via versatile manufacturing processes, namely, hot-melt coating, spray-drying, solid lipid extrusion, 3D printing, tableting, and high-pressure homogenization, among others. Dosage forms such as tablets, orally disintegrating films, multiparticulate systems, nano and microparticles, pellets, and 3D-printed forms are the result2,3,4.
LBEs possess the &#34;General Recognized as Safe&#34; status, low toxicity, good biocompatibility, and improved patient tolerance. Their natural origin and broad availability allow them to empower green and sustainable pharmaceutical manufacturing. Nevertheless, the use of LBEs has been associated with unstable dosage forms. Alterations in the properties of lipid-based products after storage have been widely reported. The solid-state of LBEs and the existence of lipid polymorphism are considered the main reasons for the instability of lipid-based dosage forms5,6,7,8.
The mechanical and physical properties of lipids are closely related to their crystallization properties and the structure of their crystal network, which shows distinct hierarchies of structural organization. When lipids are used in the manufacturing of pharmaceutical products, the crystal structure is affected by the process parameters applied, such as temperature, organic solvents, shear, and mechanical forces, which in turn affects the performance of the pharmaceutical product5,7,9,10,11,12. To understand this structure-function relationship, it is important to know the fundaments of lipid crystallization and crystal structure and analytical methods to screen them.
At the molecular level, the smallest unit of a lipid crystal is termed a &#34;unit cell.&#34; A regular three-dimensional repetition of unit cells builds the crystal lattices, with stronger molecular interactions alongside their lateral directions than the longitudinal ones, explaining the layered construction of lipid crystals. The repeated cross-sectional packing of hydrocarbon chains is known as sub-cell1,12,13 (Figure 1). Lamellae are the lateral packing of lipid molecules. In the crystal package, the interfaces between different lamellae are made of methyl end groups, whereas the polar glycerol groups are placed at the interior parts of the lamella14. To differentiate each fatty-acid chain in the lamella, the term leaflet is employed, which represents a sublayer composed of single fatty-acid chains. Acylglycerols can be arranged in double (2L) or triple (3L) leaflet chain lengths14. The surface energy of the lamellae drives them to epitaxially stack to each other, to provide nano-crystallites. Different processing factors such as cooling temperature and rate affect the number of stacked lamellae and thereby, the crystallite thickness (~10-100 nm). Aggregation of crystallites leads to the formation of spherulites in micro-scale, and the aggregation of spherulites provides the crystal network of LBEs with defined macroscopic behavior13.
Solid-state transitions start at the molecular level. The geometrical transition from one sub-cell to another is called polymorphism. Three major polymorphs of &#945;-, &#946;&#39;-, and &#946;-form are usually found in acylglycerols, ordered according to increased stability. Tilting of the lamella with respect to end-groups occurs during polymorphic transitions1,13. Storage and melt-mediated polymorphic transitions are experienced by LBEs. Storage transitions occur when the metastable form is stored below its melting temperature, whereas melt-mediated transitions happen as the temperature rises above the melting point of a metastable form provoking melting and successive crystallization of the more stable form.
Furthermore, phase separation and crystal growth can also occur. Phase separation is driven by initial multiphasic crystallization and growth of one phase or more. Particle-particle interactions, including sintering, molecular interactions, microstructural features, and foreign components, can also trigger crystal growth1,5.
Monitoring the solid-state transitions of LBEs and their impact on the performance of dosage forms is of significant importance. Among others, differential scanning calorimetry (DSC) and x-ray diffraction, specifically simultaneous small and wide-angle X-ray scattering (SWAXS), are two gold standards for assessing lipid solid-state.
DSC is commonly used to measure the enthalpy changes of the material of interest associated with the heat flow as a function of time and temperature. The method is widely used for the screening of thermal behavior of lipids, such as possible pathways of melting and crystallization, corresponding temperature and enthalpy of different polymorphic forms, as well as minor and main fractions of lipid compositions. These data can be used to depict the heterogeneity, multiple phases, and lipid polymorphism5,7,13.
X-ray diffraction techniques are the most powerful methods for structure determination in the solid state. Possessing ordered nanostructure with repeated lamellae, the reflection of x-ray beam from lipid crystals can be investigated using Bragg&#39;s law:
d = &#955;/2sin&#952;&#160; &#160; &#160;(Equation 1)
where &#955; is the x-ray wavelength of 1.542 &#197;, &#952; is the diffraction angle of the scattered beam, and d is the interplanar spacing of repeated layers, defined as lamella length in lipids. The small-angle region of the x-ray can be perfectly used to detect the long-spacing pattern and calculate the lamella length (d). The larger the repeated distance d, the smaller the scattering angle (1-15&#176;, small-angle region) since d is inversely proportional to sin &#952;. The sub-cell arrangement of lipids can be characterized as the short-spacing pattern in the wide-angle region of the x-ray diffraction. Both the long- and short-spacing patterns of lipids (lamella length and sub-cell arrangement) can be used to indicate the monotropic polymorphic transformation. For example, the &#945;-form (hexagonal) can be altered to &#946; (triclinic) due to a change in the angle of tilt of the chains, with alterations in the lamella length (long-spacing pattern, in the small-angle region, 1-15&#176;) and in the cross-sectional packing mode (short-spacing pattern, in the wide-angle region, 16-25&#176;) (Figure 2).
The information obtained from the SAXS region can further be used to investigate the crystal growth by measuring its thickness (D) via the Scherrer equation15:
D = K&#955;/FWHMcos&#952;&#160; &#160; &#160;(Equation 2)
Where, FWHM is the width in radians of the diffraction maximum measured at a half-way height between the background and the peak, generally known as full-width at half-maximum (FWHM); &#952; is the diffraction angle; &#955; is the X-ray wavelength (1.542 &#197;) and K (Scherrer constant) is a dimensionless number that provides information about the shape of the crystal (in case of the absence of detailed shape information K = 0.9 is a good approximation). Please note that the Scherrer equation can be used to estimate mean crystal sizes of up to about 100 nm since the peak broadening is inversely proportional to the crystallite size. Therefore, its application is useful for determining the thickness of nanoplatelets and, indirectly, the number of aggregated lamellae. Examples of using this well-known approach for screening the crystal properties of lipids in the pharmaceutical formulation development and the corresponding instability in product performance can be found in5,12,16,17,18.
Monitoring the solid-state of LBEs within each developmental stage through well-established analytical techniques provide an effective strategy for designing high-performance manufacturing processes and stable lipid-based pharmaceutical products.
This publication presents the critical application of a comprehensive solid-state analysis of LBEs for monitoring the changes in solid-state and its correlation to the alteration in the release profile of active pharmaceutical ingredient (API) from the pharmaceutical dosage form. Multiparticulate systems based on lipid-coated API crystals via hot-melt coating, and nano-lipid suspensions produced via high-pressure homogenization are taken as case studies. The focus of this publication is the application of powder x-ray diffraction and DSC as analytical tools. The first two examples show the effect of polymorphic transformation and crystal growth, respectively, on the alteration in API release from coated samples. The last example reveals the correlation between the stable solid-state of lipid and the stable performance of the pharmaceutical product in lipid-coated multiparticulate systems and in nano-lipid suspensions.

<table><tbody><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma-Aldrich</td><td>223506</td><td></td></tr><tr><td>Cassettes with a cellulose membrane bag with a cut-off of 7000 Da, Thermo Scientific Slide-A-Lyzer 7K</td><td>Fisher Scientific Inco, USA</td><td></td><td></td></tr><tr><td>Control software of x-ray system</td><td>HECUS dedicated house equipment</td><td></td><td></td></tr><tr><td>Control unit of x-ray system</td><td>HECUS dedicated house equipment</td><td></td><td></td></tr><tr><td>Differential scanning calorimeter (DSC) aluminum crucibles and lids</td><td>Netzsch, Germany</td><td></td><td></td></tr><tr><td>Differential scanning calorimeter DSC 204 F1 Phoenix (NETZSCH, Germany).</td><td>Netzsch, Germany</td><td></td><td></td></tr><tr><td>Dipalmitoylphosphatidylcholine (DPPC)</td><td>Sigma-Aldrich</td><td>850355P</td><td></td></tr><tr><td>Dissolution paddle apparatus II, Erweka DT 828 LH</td><td>Erweka GmbH, Langen, Germany</td><td></td><td></td></tr><tr><td>Dynasan 116</td><td>IOI OLEO</td><td></td><td>Tripalmitin</td></tr><tr><td>Geleol</td><td>Gattefosse</td><td></td><td>Glyceryl monosterarate&amp;nbsp;</td></tr><tr><td>KCl&amp;nbsp;</td><td>Sigma-Aldrich</td><td>529552</td><td></td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>P0662</td><td></td></tr><tr><td>Kolliphor P 188</td><td>BASF Chem Trade</td><td></td><td>Poloxamer 188&amp;nbsp;</td></tr><tr><td>MgCl2&amp;middot;6H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma-Aldrich</td><td>M2670</td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma-Aldrich</td><td>S9763</td><td></td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>S9888</td><td></td></tr><tr><td>Netzsch DSC 204F1 Software Version 8.0.1</td><td>Netzsch, Germany</td><td>6.239.2-64.51.00</td><td></td></tr><tr><td>Origin Pro (OriginLab, Northampton, MA) (statistical software</td><td>OriginLab, Northampton, MA</td><td></td><td></td></tr><tr><td>Proteous Analysis Software</td><td>Netzsch, Germany</td><td></td><td></td></tr><tr><td>Tween 65</td><td></td><td></td><td>Polysorbate 65</td></tr><tr><td>Witepsol PMF 1683</td><td>IOI OLEO</td><td></td><td>Triglycerol ester of stearatic/palmitic acid (partially esterified)</td></tr><tr><td>Witepsol PMF 282</td><td>IOI OLEO</td><td></td><td>Diglycerol ester of stearic acid&amp;nbsp;</td></tr><tr><td>X-ray HECUS system composed of a point-focusing camera and two linearly positioned sensitive detectors</td><td>HECUS dedicated house equipment</td><td></td><td></td></tr></tbody></table>

a package of established analytical tools to investigate the solid-state alteration of lipid-based excipients

Lipid-based excipients (LBEs) are low-toxic, biocompatible, and natural-based, and their application supports the sustainability of pharmaceutical manufacturing. However, the major challenge is their unstable solid-state, affecting the stability of the pharmaceutical product. Critical physical properties of lipids for their processing-such as melt temperature and viscosity, rheology, etc.-are related to their molecular structure and their crystallinity. Additives, as well as thermal and mechanical stress involved in the manufacturing process, affect the solid-state of lipids and thus the performance of pharmaceutical products thereof. Therefore, understanding the alteration in the solid-state is crucial. In this work, the combination of powder x-ray diffraction and differential scanning calorimetry (DSC) is introduced as the gold standard for the characterization of lipids' solid state. X-ray diffraction is the most efficient method to screen polymorphism and crystal growth. The polymorphic arrangement and the lamella length are characterized in the wide- and small-angle regions of x-ray diffraction, respectively. The small-angle x-ray scattering (SAXS) region can be further used to investigate crystal growth. Phase transition and separation can be indicated. DSC is used to screen the thermal behavior of lipids, estimate the miscibility of additives and/or active pharmaceutical ingredients (API) in the lipid matrix, and provide phase diagrams. Four case studies are presented in which LBEs are either used as a coating material or as an encapsulation matrix to provide lipid-coated multiparticulate systems and lipid nanosuspensions, respectively. The lipid solid-state and its potential alteration during storage are investigated and correlated to the alteration in the API release. Qualitative microscopical methods such as polarized light microscopy and scanning electron microscopy are complementary tools to investigate micro-level crystallization. Further analytical methods should be added based on the selected manufacturing process. The structure-function-processability relationship should be understood carefully to design robust and stable lipid-based pharmaceutical products.

Correlation between polymorphic transition of lipid and API release in lipid-coated API crystals: 
API crystals coated with glycerol monostearate are measured via DSC and x-ray directly after coating and after 3 months of storage under accelerated conditions (40 &#176;C, 75% relative humidity)7. Glyceryl monostearate is a multiphasic system containing 40%-55% monoglycerides, 30%-45% diglycerides, and 5%-15% glycerides, mainly tristearin19. The...

Watch this Scientific Journal Video about A Package of Established Analytical Tools to Investigate the Solid-State Alteration of Lipid-Based Excipients at JoVE.com

A Package of Established Analytical Tools to Investigate the Solid-State Alteration of Lipid-Based Excipients

This publication shows the application of x-ray diffraction and differential scanning calorimetry as gold standards for investigating the solid-state of lipid-based excipients (LBEs). Understanding the solid-state alteration in LBEs and its effect on the performance of pharmaceutical products thereof is the key factor for manufacturing robust lipid-based dosage forms.

Lipid-based excipients (LBEs) are low-toxic, biocompatible, and natural-based, and their application supports the sustainability of pharmaceutical ...

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Research

JoVE Journal

Chemistry

2.0K Views.  GmbH. De combinatie van differentiële scanningcalorimetrie en röntgendiffractie is een gouden standaard voor onderzochte solid-state veranderingen van lipide-gebaseerde farmaceutische producten. De gecombineerde methodologie is een sterk hulpmiddel voor het screenen van het polymorfisme, de kristalgroei en de gezichtsovergang van lipiden in combinatie met de screening, het terminale gedrag en de zichtbaarheid van het geneesmiddel met lipidenmatrix. Deze methodologie kan worden gebruikt om de inte...

Video: A Package of Established Analytical Tools to Investigate the Solid-State Alteration of Lipid-Based Excipients

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous number of imaging techniques currently available and the progress in instrumentation, there is still a need for highly sensitive probes that are suitable for in vivo imaging. One typical problem of available preclinical fluorescent probes is their rapid clearance in vivo, which reduces their imaging sensitivity. To circumvent rapid clearance, increase number of dye molecules at the target site, and thereby reduce background autofluorescence, encapsulation of the near-infrared fluorescent dye, DY-676-COOH in liposomes and verification of its potential for in vivo imaging of inflammation was done. DY-676 is known for its ability to self-quench at high concentrations. We first determined the concentration suitable for self-quenching, and then encapsulated this quenching concentration into the aqueous interior of PEGylated liposomes. To substantiate the quenching and activation potential of the liposomes we use a harsh freezing method which leads to damage of liposomal membranes without affecting the encapsulated dye. The liposomes characterized by a high level of fluorescence quenching were termed Lip-Q. We show by experiments with different cell lines that uptake of Lip-Q is predominantly by phagocytosis which in turn enabled the characterization of its potential as a tool for in vivo imaging of inflammation in mice models. Furthermore, we use a zymosan-induced edema model in mice to substantiate the potential of Lip-Q in optical imaging of inflammation in vivo. Considering possible uptake due to inflammation-induced enhanced permeability and retention (EPR) effect, an always-on liposome formulation with low, non-quenched concentration of DY-676-COOH (termed Lip-dQ) and the free DY-676-COOH were compared with Lip-Q in animal trials.

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

The use of fluorophores for in vivo imaging can be greatly limited by opsonization, rapid clearance, low detection sensitivity and cytotoxic effects on the host. Encapsulation of fluorophores in liposomes by film hydration and extrusion leads to fluorescence quenching and protection which enables in vivo imaging with high detection sensitivity.

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous ...

Bioengineering

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient magnification to examine membrane substructures and even individual molecules. AFM can act as a force probe to measure interactions and mechanical properties of membranes. Supported lipid bilayers are conventionally used as membrane models in AFM studies. In this protocol, we demonstrate how to prepare supported bilayers and characterize their structure and mechanical properties using AFM. These include bilayer thickness and breakthrough force. 
The information provided by AFM imaging and force spectroscopy help define mechanical and chemical properties of membranes. These properties play an important role in cellular processes such as maintaining cell hemostasis from environmental stress, bringing membrane proteins together, and stabilizing protein complexes.

We describe a protocol for preparation of supported lipid bilayers and its characterization using atomic force microscopy and force spectroscopy.

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient ...

Gramicidin-based Fluorescence Assay; for Determining Small Molecules Potential for Modifying Lipid Bilayer Properties

Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter lipid bilayer properties. This is important because membrane proteins are energetically coupled to their host bilayer by hydrophobic interactions. Changes in bilayer properties thus alter membrane protein function, which provides an indirect way for amphiphiles to modulate protein function and a possible mechanism for "off-target" drug effects. We have previously developed an electrophysiological assay for detecting changes in lipid bilayer properties using linear gramicidin channels as probes 3,12. Gramicidin channels are mini-proteins formed by the transbilayer dimerization of two non-conducting subunits. They are sensitive to changes in their membrane environment, which makes them powerful probes for monitoring changes in lipid bilayer properties as sensed by bilayer spanning proteins. We now demonstrate a fluorescence assay for detecting changes in bilayer properties using the same channels as probes. The assay is based on measuring the time-course of fluorescence quenching from fluorophore-loaded large unilamellar vesicles due to the entry of a quencher through the gramicidin channels. We use the fluorescence indicator/quencher pair 8-aminonaphthalene-1,3,6-trisulfonate (ANTS)/Tl+ that has been successfully used in other fluorescence quenching assays 5,13. Tl+ permeates the lipid bilayer slowly 8 but passes readily through conducting gramicidin channels 1,14. The method is scalable and suitable for both mechanistic studies and high-throughput screening of small molecules for bilayer-perturbing, and potential "off-target", effects. We find that results using this method are in good agreement with previous electrophysiological results 12.

We introduce a fast fluorescence-based assay that monitors the rate of fluorescence quenching as a measure of gramicidin channel activity. The gramicidin channels are used as molecular force transducers to monitor changes in lipid bilayer properties as sensed by bilayer spanning proteins.

Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter ...

Biology

Spontaneous Formation and Rearrangement of Artificial Lipid Nanotube Networks as a Bottom-Up Model for Endoplasmic Reticulum

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense lipidic nanotubes that are, in terms of morphology and dynamics, reminiscent of ER. The networks are derived from phospholipid double bilayer membrane patches adhering to a transparent Al2O3 substrate. The adhesion is mediated by Ca2+ in the ambient buffer. Subsequent depletion of Ca2+ by means of BAPTA/EDTA causes retraction of the membrane, resulting in spontaneous lipid nanotube network formation. The method only comprises phospholipids and microfabricated surfaces for simple formation of an ER model and does not require the addition of proteins or chemical energy (e.g., GTP or ATP). In contrast to the 3D morphology of the cellular endoplasmic reticulum, the model is two-dimensional (albeit the nanotube dimensions, geometry, structure, and dynamics are maintained). This unique in vitro ER model consists of only a few components, is easy to construct, and can be observed under a light microscope. The resulting structure can be further decorated for additional functionality, such as the addition of ER-associated proteins or particles to study transport phenomena among the tubes. The artificial networks described here are suitable structural models for the cellular ER, whose unique characteristic morphology has been shown to be related to its biological function, whereas details regarding formation of the tubular domain and rearrangements within are still not completely understood. We note that this method uses Al2O3 thin-film-coated microscopy coverslips, which are commercially available but require special orders. Therefore, it is advisable to have access to a microfabrication facility for preparation.

Solid-supported, protein-free, double phospholipid bilayer membranes (DLBM) can be transformed into complex and dynamic lipid nanotube networks and can serve as 2D bottom-up models of the endoplasmic reticulum.

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense ...

Preparation and Characterization of Nanoliposomes for the Entrapment of Bioactive Hydrophilic Globular Proteins

Liposome nanocapsules have been applied for many purposes in the pharmaceutical, cosmetic, and food industries. Attributes of liposomes include their biocompatibility, biodegradability, non-immunogenicity, non-toxicity, and ability to entrap both hydrophilic and hydrophobic compounds. The classical hydration of thin lipid films in an organic solvent is applied herein as a technique to encapsulate tarin, a plant lectin, in nanoliposomes. Nanoliposome size, stability, entrapment efficiency, and morphological characterization are described in detail. The nanoliposomes are prepared using 1,2-dioleoyl-sn-glycerol-3-phosphoethanolamine (DOPE), 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[amino(polyethylene glycol)-2000] (ammonium salt; DSPE-MPEG 2000), and cholesterylhemisuccinate (CHEMS) as the main constituents. Lipids are first dissolved in chloroform to obtain a thin lipid film that is subsequently rehydrated in ammonium sulfate solution containing the protein to be entrapped and incubated overnight. Then, sonication and extrusion techniques are applied to generate nanosized unilamellar vesicles. The size and polydispersity index of the nanovesicles are determined by dynamic light scattering, while nanovesicle morphology is assessed by scanning electron microscopy. Entrapment efficiency is determined by the ratio of the amount of unencapsulated protein to original amount of initially loaded protein. Homogeneous liposomes are obtained with an average size of 155 nm and polydispersity index value of 0.168. A high entrapment efficiency of 83% is achieved.

This study describes classical hydration using the thin lipid film method for nanoliposome preparation followed by nanoparticle characterization. A 47 kDa-hydrophilic and globular protein, tarin, is successfully encapsulated as a strategy to improve stability, avoid fast clearance, and promote controlled release. The method can be adapted to hydrophobic molecules encapsulation.

Liposome nanocapsules have been applied for many purposes in the pharmaceutical, cosmetic, and food industries. Attributes of liposomes include their ...

A Quantitative Fluorescence Microscopy-based Single Liposome Assay for Detecting the Compositional Inhomogeneity Between Individual Liposomes

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes all liposomes of the ensemble to be identical. However, new experimental platforms able to observe liposomes at the single-particle level have made it possible to perform highly sophisticated and quantitative studies on protein-membrane interactions or drug carrier properties on individual liposomes, thus avoiding errors from ensemble averaging. Here we present a protocol for preparing, detecting, and analyzing single liposomes using a fluorescence-based microscopy assay, facilitating such single-particle measurements. The setup allows for imaging individual liposomes in a massive parallel manner and is employed to reveal intra-sample size and compositional inhomogeneities. Additionally, the protocol describes the advantages of studying liposomes at the single liposome level, the limitations of the assay, and the important features to be considered when modifying it to study other research questions.

This protocol describes the fabrication of liposomes and how these can be immobilized on a surface and imaged individually in a massive parallel manner using fluorescence microscopy. This allows for the quantification of the size and compositional inhomogeneity between single liposomes of the population.

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes ...

Microfluidic Production of Lysolipid-Containing Temperature-Sensitive Liposomes

The presented protocol enables a high-throughput continuous preparation of low temperature-sensitive liposomes (LTSLs), which are capable of loading chemotherapeutic drugs, such as doxorubicin (DOX). To achieve this, an ethanolic lipid mixture and ammonium sulfate solution are injected into a staggered herringbone micromixer (SHM) microfluidic device. The solutions are rapidly mixed by the SHM, providing a homogeneous solvent environment for liposomes self-assembly. Collected liposomes are first annealed, then dialyzed to remove residual ethanol. An ammonium sulfate pH-gradient is established through buffer exchange of the external solution by using size exclusion chromatography. DOX is then remotely loaded into the liposomes with high encapsulation efficiency (&#62; 80%). The liposomes obtained are homogenous in size with Z-average diameter of 100 nm. They are capable of temperature-triggered burst release of encapsulated DOX in the presence of mild hyperthermia (42 &#176;C). Indocyanine green (ICG) can also be co-loaded into the liposomes for near-infrared laser-triggered DOX release. The microfluidic approach ensures high-throughput, reproducible and scalable preparation of LTSLs.

The protocol presents the optimized parameters for preparing thermosensitive liposomes using the staggered herringbone micromixer microfluidics device. This also allows co-encapsulation of doxorubicin and indocyanine green into the liposomes and the photothermal-triggered release of doxorubicin for controlled/triggered drug release.

The presented protocol enables a high-throughput continuous preparation of low temperature-sensitive liposomes (LTSLs), which are capable of loading ...

Preparation, Administration, and Assessment of In Vivo Tissue-Specific Cellular Uptake of Fluorescent Dye-Labeled Liposomes

There is a growing interest in using liposomes to deliver compounds in vivo particularly for targeted treatment approaches. Depending on the liposome formulation, liposomes may be preferentially taken up by different cell types in the body. This may influence the efficacy of the therapeutic particle as progression of different diseases is tissue- and cell-type-specific. In this protocol, we present one method for synthesizing and fluorescently labeling liposomes using DSPC, cholesterol, and PEG-2000 DSPE and the lipid dye DiD as a fluorescent label. This protocol also presents an approach for delivering liposomes in vivo and assessing cell-specific uptake of liposomes using flow cytometry. This approach can be used to determine the types of cells that take up liposomes and quantify the distribution and proportion of liposome-uptake across cell types and tissues. While not mentioned in this protocol, additional assays such as immunofluorescence and single-cell fluorescence imaging on a cytometer will strengthen any findings or conclusions made as they permit assessment of intracellular staining. Protocols may also need to be adapted depending on the tissue(s) of interest.

The goal of this protocol is to synthesize fluorescently-labeled liposomes and use flow cytometry to identify in vivo localization of liposomes at a cellular level.

There is a growing interest in using liposomes to deliver compounds in vivo particularly for targeted treatment approaches. Depending on the liposome ...

Examining BCL-2 Family Function with Large Unilamellar Vesicles

The BCL-2 (B cell CLL/Lymphoma) family is comprised of approximately twenty proteins that collaborate to either maintain cell survival or initiate apoptosis1. Following cellular stress (e.g., DNA damage), the pro-apoptotic BCL-2 family effectors BAK (BCL-2 antagonistic killer 1) and/or BAX (BCL-2 associated X protein) become activated and compromise the integrity of the outer mitochondrial membrane (OMM), though the process referred to as mitochondrial outer membrane permeabilization (MOMP)1. After MOMP occurs, pro-apoptotic proteins (e.g., cytochrome c) gain access to the cytoplasm, promote caspase activation, and apoptosis rapidly ensues2.
In order for BAK/BAX to induce MOMP, they require transient interactions with members of another pro-apoptotic subset of the BCL-2 family, the BCL-2 homology domain 3 (BH3)-only proteins, such as BID (BH3-interacting domain agonist)3-6. Anti-apoptotic BCL-2 family proteins (e.g., BCL-2 related gene, long isoform, BCL-xL; myeloid cell leukemia 1, MCL-1) regulate cellular survival by tightly controlling the interactions between BAK/BAX and the BH3-only proteins capable of directly inducing BAK/BAX activation7,8. In addition, anti-apoptotic BCL-2 protein availability is also dictated by sensitizer/de-repressor BH3-only proteins, such as BAD (BCL-2 antagonist of cell death) or PUMA (p53 upregulated modulator of apoptosis), which bind and inhibit anti-apoptotic members7,9. As most of the anti-apoptotic BCL-2 repertoire is localized to the OMM, the cellular decision to maintain survival or induce MOMP is dictated by multiple BCL-2 family interactions at this membrane.
 Large unilamellar vesicles (LUVs) are a biochemical model to explore relationships between BCL-2 family interactions and membrane permeabilization10. LUVs are comprised of defined lipids that are assembled in ratios identified in lipid composition studies from solvent extracted Xenopus mitochondria (46.5% phosphatidylcholine, 28.5% phosphatidylethanoloamine, 9% phosphatidylinositol, 9% phosphatidylserine, and 7% cardiolipin)10. This is a convenient model system to directly explore BCL-2 family function because the protein and lipid components are completely defined and tractable, which is not always the case with primary mitochondria. While cardiolipin is not usually this high throughout the OMM, this model does faithfully mimic the OMM to promote BCL-2 family function. Furthermore, a more recent modification of the above protocol allows for kinetic analyses of protein interactions and real-time measurements of membrane permeabilization, which is based on LUVs containing a polyanionic dye (ANTS: 8-aminonaphthalene-1,3,6-trisulfonic acid) and cationic quencher (DPX: p-xylene-bis-pyridinium bromide)11. As the LUVs permeabilize, ANTS and DPX diffuse apart, and a gain in fluorescence is detected. Here, commonly used recombinant BCL-2 family protein combinations and controls using the LUVs containing ANTS/DPX are described.

Biochemically-defined large unilamellar vesicles (LUVs) are a convenient model system to analyze BCL-2 family interactions with immediate implications in better understanding the mitochondrial pathway of apoptosis. A method to produce LUVs, along with standard BCL-2 family protein combinations and controls to examine LUV permeabilization, are presented.

The BCL-2 (B cell CLL/Lymphoma) family is comprised of approximately twenty proteins that collaborate to either maintain cell survival or initiate ...

Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify and explore the function of these important proteins. However, classical preparations, such as planar bilayers, limit the manipulations and experiments that can be performed on the reconstituted channel and its membrane environment. The more cell-like structure of giant liposomes permits traditional patch-clamp experiments without sacrificing control of the lipid environment.
Electroformation is an efficient mean to produce giant liposomes &gt;10 &mu;m in diameter which relies on the application of alternating voltage to a thin, ordered lipid film deposited on an electrode surface. However, since the classical protocol calls for the lipids to be deposited from organic solvents, it is not compatible with less robust membrane proteins like ion channels and must be modified. Recently, protocols have been developed to electroform giant liposomes from partially dehydrated small liposomes, which we have adapted to protein-containing liposomes in our laboratory.
We present here the background, equipment, techniques, and pitfalls of electroformation of giant liposomes from small liposome dispersions. We begin with the classic protocol, which should be mastered first before attempting the more challenging protocols that follow. We demonstrate the process of controlled partial dehydration of small liposomes using vapor equilibrium with saturated salt solutions. Finally, we demonstrate the process of electroformation itself. We will describe simple, inexpensive equipment that can be made in-house to produce high-quality liposomes, and describe visual inspection of the preparation at each stage to ensure the best results.

Reconstituting functional membrane proteins into giant liposomes of defined composition is a powerful approach when combined with patch-clamp electrophysiology. However, conventional giant liposome production may be incompatible with protein stability. We describe protocols for producing giant liposomes from pure lipids or small liposomes containing ion channels.

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify ...

A Package of Established Analytical Tools to Investigate the Solid-State Alteration of Lipid-Based Excipients

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