Results
The reduction of the RSD of the measured API concentrations shows how the mixture approaches homogeneity with rising mixing
times (see Figure 1). A time of 30 min was chosen as sufficient to view the mixture of DC-mannitol with micronized ascorbic acid as homogeneous
(RSD = 0.67%). The mixing behavior of a blend is dependent on the API and the excipient, as well as on mixer type, scale,
and the degree of filling of the mixer. As the latter parameters were constant for all assessed blends, differences in homogeneity
must be due to either the API or the excipient. In this case, the micronized hydrophobic particles of riboflavin tend to re-agglomerate
during mixing. This re-agglomeration is why at first the homogeneity decreases before the mixture reaches a steady state (see
Figure 1).
The resulting mixing time of 30 min seems to be rather high. It has to be taken into account that this small laboratory-scale
mixing unit is certainly not optimized. More importantly, however, the micronized API granules have a tendency of agglomeration
to each other due to their high surface energy. This binding force has to be broken up and replaced by an alternative binding
force—adsorption and van der Vaals interaction–with the carrier surface. This is a dynamic equilibrium process and takes more
time than just a statistical distribution of different particles in space.
Figure 3: Comparison of the surface area and pore volumes of different excipients available for direct compression. DC is
direct compression. Parkteck M 200 is a proprietary product of DC-grade mannitol (Merck KGaA). Sp is specific.
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The comparison of different DC-mannitols at optimum mixing time reveals differences in the homogeneity of such a mixture with
micronized ascorbic acid and riboflavin (see Figure 2). Clearly, for a hydrophilic API, the achievable homogeneity is greater than for a hydrophobic API. In this case, the different
attraction forces of the hydrophobic API to a hydrophilic carrier cause more API particles to re-agglomerate rather than bind
to the carrier surface. This is not a surprising observation because this relationship would be true for all excipients. There
are, however, differences in the achievable blend uniformity among the compared carriers. The best homogeneity for both API
cases was found for the excipient with the highest surface area, DC-Mannitol M (see Table I). This observation gives a hint for a correlation of BET surface area and/or pore volume to the achievable homogeneity. There
also are significant differences between spray-dried and granulated DC-mannitol even having similar BET-surfaces (see Figure 3). The quality of the surface structure, not only the quantitative size of the surface area, seems to be relevant.
To prove an adsorption of the API to the excipient surface with a certain force, the remaining concentration of ascorbic acid
and riboflavin was measured after 15 min on an air-jet sieve. By this procedure, a separation of fine API particles from the
carrier can be expected if they were not strongly adsorbed. A recovery of 100% would mean a perfectly strong adsorption of
the API to the carrier while a recovery of 0% shows no absorption to the carrier.
Figure 4: Comparison of the API concentration measured after 15 min in an air-jet sieve using either granulated direct-compression
(DC)-Mannitol B, spray-dried DC-Mannitol A, or DC-mannitol M as excipients for the model drugs of ascorbic acid or riboflavin.
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A much stronger adsorption was found for the spray-dried DC-mannitols in comparison to the granulated quality (see Figure 4). For low API concentrations of a hydrophilic drug, both spray-dried mannitols show similar results. Using higher API loads,
it was demonstrated that the higher surface area of DC-Mannitol M shows advantages of a higher binding capacity. This effect
was confirmed with a hydrophobic API, riboflavin. This finding may again result from the different surface structure of the
investigated excipients. The lower recovery of hydrophobic API again confirms a weaker force of surface adsorption by this
class of API.
Figure 5: Scanning electron microscope (SEM) image showing a mixture of direct-compression (DC)-mannitol M 200 and micronized
ascorbic acid (drug load 1% w/w).
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To visualize the API distribution on the excipients' surface, a scanning electron microscope (SEM) was employed. Figure 5 shows the SEM image of a mixture of ascorbic acid and DC-mannitol M. The micronized API particles are readily identifiable
due to the different crystal structures of API and carrier (colorization performed manually). The API crystals were found
within the pore structure of the much larger excipient particles. Figure 6 shows the SEM image of spray-dried DC-Mannitol A and ascorbic acid. In this case, less areas are present that are suitable
for the absorption of the API. The overall surface is less structured. A similar distribution on the excipients' surface was
determined for the hydrophobic model drug riboflavin (see Figure 7).
Figure 6: Scanning electron microscope (SEM) image showing a mixture of spray-dried direct compression (DC)-Mannitol A and
micronized ascorbic acid (drug load 1% w/w).
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The importance of the surface area and the pore volume of an excipient for the homogeneity of the mixture was demonstrated.
In the next step, the surface area and porosity of various excipients available for direct compression were analyzed using
the BET method (nitrogen adsorption). As the API is adsorbed to a porous surface, the observed differences of the excipients
may give rise to a different behavior in the adsorption of the micronized APIs (see Figure 3).
Figure 7: Mixture of direct-compression (DC)- mannitol M 200 with micronized riboflavin (drug load 1% w/w). Light microscope
with 40 x magnification. The yellow particles of the API are clearly visible in the porosity of the carrier surface.
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This study showed that stable mixtures of much smaller micronized API particles with DC-excipients can be achieved. The next
question examined was whether this approach was suitable for the DC process for an actual formulation.
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